Thesis
Investigating the subsurface biosphere of a hypersaline environment - the Dead Sea (Levant)
THOMAS, Camille
Abstract
In the framework of the Dead Sea Deep Drilling Project, a geomicrobiological investigation has taken place to understand the extent and characteristics of life in the hypersaline sediment of the Dead Sea. The DNA recovered in the ICDP cores suggests that different microbial assemblages are associated with particular sedimentary facies, regardless of their depth and in situ salinity. Since this facies are controlled by changes in the evaporation-precipitation ratio in the region, our data suggest that subsurface microbial assemblages are themselves highly influenced by climatic variations at the time of sedimentation. In particular, humid periods allow the development of varied metabolisms such as sulfate reduction, methanogenesis and potentially anaerobic methane oxidation, deeply influencing the carbon and sulfur cycles of the lake, and subsequently allowing the formation of diagenetic Fe-S minerals. These results reveal the importance of considering microbial impact on archives retrieved from lacustrine drilling project, even in extreme environments.
Reference
THOMAS, Camille. Investigating the subsurface biosphere of a hypersaline environment - the Dead Sea (Levant). Thèse de doctorat : Univ. Genève, 2015, no. Sc. 4769
URN : urn:nbn:ch:unige-737244 DOI : 10.13097/archive-ouverte/unige:73724
Available at: http://archive-ouverte.unige.ch/unige:73724
Disclaimer: layout of this document may differ from the published version.
1 / 1 UNIVERSITE DE GENEVE FACULTE DES SCIENCES Département des sciences de la Terre Professeur D. Ariztegui
Investigating the Subsurface Biosphere of a Hypersaline Environment – The Dead Sea (Levant)
THESE
présentée à la Faculté des sciences de l’Université de Genève pour obtenir le grade de Docteur ès sciences, mention sciences de la Terre
par Camille Thomas de Dijon (France)
Thèse N° 4769
GENEVE Atelier de reprographie ReproMail 2015
This thesis was accomplished with the financial support of the Swiss National Science Foundation through grants 200021-132529 and 200020-149221/1 at the Earth Science Department of the University of Geneva, Geneva, Switzerland
Remerciements
S’il est vrai que la thèse peut parfois décourager un doctorant du monde de la recherche, j’ai peur que pour moi elle ne l’ait rendu difficilement remplaçable. Les opportunités qui m’ont été offertes, les rencontres que j’y ai faites et la confiance qui m’a été accordée ont certainement contribué à faire de moi ce que je suis aujourd’hui et je serai toujours reconnaissant envers ces personnes qui m’ont accompagné pendant ces quatre années. En particulier, je remercie mon directeur de thèse, qui m’a fait confiance, qui m’a toujours incité à étendre mes connaissances, a soutenu chacune de mes initiatives et a su me tirer vers le haut et me redonner confiance lors des moments de doute et de remise en question. Daniel est un scientifique formidable, un superviseur disponible et attentif, un collègue altruiste et juste, bref un directeur de thèse exceptionnel. Sans lui, cette thèse serait bien évidemment différente, et moi-même je le serais. Merci Daniel. Je tiens aussi à remercier mon collègue et collaborateur Danny Ionescu. Danny taught me a lot, chaperoned me in the field of microbial ecology and shared a lot with me. He also involved much of his time in the writing of the articles, and I am grateful of all he did for me. I hope to continue working with him. And of course, I thank Mina, whom I know was always present in the background when Danny was reviewing my work on Sunday nights… I learnt a lot during this PhD, I thus want to thank all the people who shared their knowledge in the lab with me, from Emanuela Reo, Franck Lejzerowicz, Laure Apothéloz-Perret-Gentil, Jan Pawlowski, to Stefan “Zwiebel” Thiele, Muriel Pacton, Arnauld Vinçon-Laugier et bien sûr Vincent Grossi, qui m’a accompagné lors de mon master, et avec qui j’ai toujours plaisir à travailler et (essayer d’) apprendre. Je remercie aussi ceux qui m’ont aidé un jour aiguillé ou conseillé, je pense notamment à Christophe Dupraz, à Robert Moritz, à Pilar Junier, à Kurt Hanselmann, ainsi qu’à Véronique Gardien, dont les recommandations ont certainement contribuées à mon engagement dans cette thèse. Merci également à Jean-Michel Jaquet et Christos Kanellopoulos pour leur confiance. Je remercie aussi tous les gens dont j’ai croisé la route au cours de ces années et avec qui j’ai partagé sciences, rires et bières. Notamment les étudiants de l’université d’Osnabrück, les compagnons microbiologistes et pédologues de Neuchâtel et Lausanne, les vacanciers géologues abonnés aux cours CUSO… I also wish to thank my colleagues remotely or closely linked to the Dead Sea project. Those that helped logistically and humanly to the core opening parties, namely Stefi, Florian, Romain… Also those that contributed to great scientific and social exchanges, Elan, Gilad, Orit, Nicolas, Elisa, Yossi, Ittai, Moti, Adi, Yael E and Yael K, Ron, Daniel, Norbert, Achim, Markus, and of course Ina. Je remercie également les membres de l’unige: Fred, François, Jacqueline, Elisabeth, Christine, Elias, Rossana, Guy, Sébastien, Georges et Eric, pour leur gentillesse et leur disponibilité. Merci à la team Corse, ceux qui ont partagés la dureté du sol sous la tente, le froid des nuits Cortenaise, les griffures du maquis, la brulure du soleil printanier, la houle méditerranéenne, les vents hurlants de Nonza, la faune agressive, la flore
i cracheuse. Antonio, Elme, et bien sûr Chloé vous étiez là et vous avez survécu. Merci d’avoir tenu le coup et d’avoir fait de ces moments d’adversité des souvenirs impérissables. Merci aussi à Monique maman locale et réconfort incarné. Enfin merci à Mario également pour tous ces moments, pour sa bonté, son sourire, tout ce que tu transmets aux étudiants et personnes qui t’entourent. J’en viens à la longue liste des collègues et amis de l’université de Genève, qui ont fait de ces années de thèse des moments inoubliables, autant dans l’adversité que dans la fête. Mes collègues du hip-hop bureau Jay Rag et SmArTy MaRtY qui ont amené et maintenu le groove, la vibe et le swag au bureau. Je remercie aussi Aurèle, pour le temps qu’il ma consacré, les connaissances (innombrables) qu’il a voulu partager avec moi, et les grands moments passés ensemble, au labo, dans le bureau, sur le terrain et en vacances ! Merci à mes amies et (gossip) girls préférées, Lina, Mélanie, Cristina, Katrina. Vous me manquez. Merci à leur pendant masculin Camille, toujours prêt pour partager les meilleures nouvelles scientifiques et people du moment. A celui qui a partagé ma vie, qui m’a nourri et blanchi pendant deux années. Merci Valentin pour tout ! Merci à Stéphanie Girardclos pour ses conseils, ses grandes discussions (son whisky). Merci à mon équipier gagnant Cyril Chelle-Michou. Merci à notre adversaire Nicolas Saintilan. La victoire est belle car l’adversaire est beau. Aux surhommes géologiques, Arnoud, Gabriel, Aymeric. A la squadra azzurra Federico, Eduardo, et à celle qui gagne, Haseeb, Mortaza. Merci Agathe pour les tête-à-tête à balayage électronique. Merci Noel, Hosseini, Chen Chen, merci les frenchies Vincent, Bertrand, merci les bahamiennes Erika, Jennifer, merci Tiago, merci Neda, merci Paula, merci les petits nouveaux Luis et Ines. Merci les maraichers, Alexandra, Roelant, Sylvain, Jerôme, Matar, Chadia et tous ceux que j’oublie. J’en rajoute encore, Laure, Stéphane, Julien, Marie, Camille, Basti, Chia-i, Hervé, Alexia, Maria, Christian. Merci aussi aux étudiants, qui ont contribué à mon bien être ici. Si j’ai envie de continuer aujourd’hui, c’est aussi parce qu’ils sont géniaux. Merci aux volées corses, aux membres de l’AEST (Antoine en particulier), et à tous ceux qui ont participé de près ou de loin à la vie maraichère : la Yourghurteria, le Volt, le Lys, le Trulli, le MacSorleys, la Ferblanterie, Kendrick, Frank, Jonsi, Thom, Beth and Geoff, tous. Il y a quelques personnes clés qui ont contribué à ce que j’ai pu entreprendre dans la géologie. Merci Lydie Prieur, pour m’avoir conseillé puis soutenu. Je suis fier aujourd’hui d’avoir été votre étudiant, et de pouvoir vous montrer quelques résultats géologiques ET biologiques. I also wish to thank Salik and Minik Rosing, who have somehow, changed the course of my short geology career. Enfin merci à Fabrice Cordey et Nicolas Olivier, qui m’ont fait confiance pendant mon master et qui ont contribué à mon entrée dans la recherche. Leur patience, leurs conseils et leur soutien m’ont permis d’entreprendre cette thèse avec confiance et optimisme. J’en viens au privé. Merci à tous mes amis, ceux de Lyon, d’Islande, du Danemark et surtout ceux de Dijon. Mes meilleurs amis à qui je n’ai pas pu consacrer beaucoup de temps au milieu de tout cela. J’ai aussi pu faire cette thèse car je sais que vous êtes à mes côtés. En particulier, merci Antoine, merci Caroline, merci Fabien, merci Mathilde C., merci Mathilde S., merci Maxime, merci Olivier.
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Je remercie aussi toute ma famille, qui m’a toujours soutenu. En particulier, merci à ma mère et mon père de croire en moi, et d’être fantastiques. Merci également à mes sœurs Juliette et Léa pour qui je n’ai pas beaucoup été là ces derniers temps, mais qui m’aident et m’équilibrent. J’espère leur apporter autant que ce qu’elles me donnent. Enfin, je terminerai par celle à qui j’ai certainement volé le plus de temps, et celle qui a dû faire plus de sacrifices que tous les autres. Merci Marie de m’avoir soutenu, merci de m’avoir accompagné, et merci pour ce que tu amènes dans ma vie. Sans toi je n’aurais pu accomplir tout ça. Je t’en remercie profondément, et suis impatient d’aborder la suite avec toi.
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Abstract
The Dead Sea Deep Drilling Project is an ICDP sponsored project that aims to reconstruct the Quaternary paleoenvironments of the Dead Sea Basin. Within this framework, a geomicrobiological investigation has been realized in order to assess the potential effect of microbial communities on the sediment, as well as their extent. Similarly to well- established studies in the oceanic subsurface, geomicrobiological analysis are now employed in lacustrine subsurface, an environment that is allegedly more susceptible to environmental and climatic disturbance than its marine counterpart. In the Dead Sea, the study also aims at qualifying the development of microbial life in extreme conditions of salinity.
A 457 m-long sedimentary core has been retrieved from the middle of the Dead Sea, in winter 2010-2011. Contrasting climatic intervals were highlighted by distinctive lithological facies such as laminated aragonitic mud for humid periods and glacial stages and halite-gypsum deposits for interglacial intervals dominated by high aridity. For the first time a deep subsurface life could be evidenced in the Dead Sea sediments through its DNA. Although living conditions are extremely harsh in this environment, a specific population has adapted and survives at depths down to at least 90 m below the present water-sediment interface, and probably deeper (200 m). Different lithological types hold specific microbial populations representative of changing environments. Members of bacterial KB1 and MSBL1 Candidate Divisions could take advantage of the osmotic solutes such as glycine betaine, available in the aragonitic sediments. Halite and gypsum sediments are dominated by extreme halophilic Archaea of the Halobacteriaceae family, which are today the only inhabitants of the lake water column. The similarity shared by gypsum-halite samples communities, and the importance of freshwater inputs for assemblages of the aragonitic sediment imply that salinity is not the only parameter influencing microbial development, but that the whole lake water balance at time of sedimentation is in part responsible for the current distribution of microbes in the sedimentary column.
Within the two main sedimentary facies previously established, archaeal metabolisms were deeply investigated using high-throughput DNA sequencing. We show that the communities are well adapted to the peculiar environment of the Dead Sea subsurface. They are able to deal with osmotic pressure using high- and low-salt-in strategies, and can also cope with high concentrations of heavy metals. Methanogenesis (from methylated compounds), for the first time identified in the Dead Sea, is an important metabolism in the aragonite sediment. Fermentation of organic matter, probably performed by some members of the Halobacteria class is common to both aragonite and gypsum-rich sediments. Genes associated with sulfur reduction have also been revealed and are associated in the sediment with microbial EPS degradation and Fe-S mineralization as revealed by SEM imaging.
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Through comparison with an active microbial mat of the Dead Sea shore, keys towards the recognition of Fe-S mineralizations of biological origin have been acquired, emphasizing the influence of a microbial sulfur cycle on mineral and organic matter. The occurrence of EPS is interpreted as an indicator of the activity of microbes in the sediment. Degradation of EPS and that of allochthonous organic matter by putative dissimilatory sulfate reduction releases H2S, enabling the formation of Fe-S precursors, which can form pyrite spherulites. Mineralizations are however small and rarely complete, supporting an incomplete sulfate reduction pathway for microbes in the Dead Sea sediment. Analogously, nanoglobules occurrence and their tight association with aragonite needles also support the use of EPS as a matrix for biologically-influenced mineralization of calcium carbonates. These results have strong implications on the fate of organic matter and on processes involved in mineralization and porosity development in the perspective of diagenetic evolution of hypersaline subsurface systems.
Higher concentration of organomineralization under the form of euhedral pyrite suggests an increase sulfate reduction activity in particular in one precise interval of the Dead Sea core. In this specific early Holocene interval, combined use of elemental and isotopic profiles of the pore water chemistry, high-resolution lithological description and lipid analysis indicate a biological signature during periods when rain dominated over evaporation. This interval also comprises a set of lipidic biomarkers indicative of anaerobic oxidation of methane. Among others, the presence of hydroxyarchaeol, pentamethyleicosene and extended archaeol highlights the reworking of biogenic methane by potential ANME Archaea. Anaerobic oxidation of methane associated with bacterial sulfate reduction is thus suggested to have disturbed organic proxies in this early Holocene interval.
In conclusion, as life extent is now better constrained in the Dead Sea subsurface, influence of the environment over its development is clear. Hypersalinity selects for hyper-halophilic groups. Additionally, humid episodes allow for development of diverse metabolisms in the water column, the latter being reflected in today’s subsurface communities. Such impact shows the need for routinely applying geomicrobiological investigations in the lacustrine realm, and of unifying methods for such research in the context of International Continental Drilling Projects. Microbial effects on mineral precipitation, isotope fractionation and organic matter preservation clearly exist in extreme environments like the Dead Sea and should always be taken into account for further paleoenvironmental reconstruction and future ecological and climatic models.
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Le Projet DSDDP (projet de forage profond de la Mer Morte), cofinancé par le consortiumRé sumé international pour les forages continentaux (ICDP), a pour objectif de reconstruire les paléo-environnements quaternaires du bassin de la Mer Morte, au Proche-Orient. Au sein de ce projet, une étude des interactions entre minéral et vivant a été entreprise afin de contraindre la potentielle influence des microbes sur leur environnement. De façon similaire à ce qui a été entrepris depuis plusieurs décennies maintenant au sein du consortium international pour les forages océaniques (IODP), des études géomicrobiogiques sont maintenant menées dans les environnements lacustres. Ces derniers sont d’ailleurs plus susceptibles aux variations environnementales et climatiques que le fond des océans. Dans le cas de la Mer Morte, l’extrême salinité de cet environnement présente également un grand intérêt quant à la compréhension des limites du vivant.
Une carotte sédimentaire de plus de 450 m de long a été forée au milieu de la Mer Morte en hiver 2010-2011. Son étude montre que les variations sédimentologiques reflètent les contrastes climatiques des 230 000 dernières années. Un facies de lamines aragonitiques marque les périodes relativement humides, caractéristiques des stades glaciaires, tandis que des unités de gypse et halite caractérisent l’augmentation de l’aridité pendant les périodes interglaciaires. Pour la première fois, la présence de vie profonde au sein du sédiment de la Mer Morte a été mise en évidence à travers son ADN. Bien que les conditions de vie soient extrêmes, des communautés microbiennes spécifiquement adaptées survivent jusqu’à des profondeurs de 90 m dans le sédiment, et s’étendent possiblement jusqu’à 200 m de profondeur. Ces communautés diffèrent en fonction de la lithologie du sédiment. Des membres des Divisions Candidates bactériennes KB1 et MSBL1 profitent des solutés osmotiques potentiellement disponibles dans le sédiment aragonitique. La classe d’Archaea hyper-halophile Halobacteriaceae, aujourd’hui principale représentante dans l’eau du lac, domine quant à elle les niveaux de gypse et halite. Les similarités d’assemblages partagées par les niveaux de gypse et halite, et l’importance des apports d’eau douce pour les communautés présentes dans les niveaux aragonitiques suggèrent que les conditions hydrologiques et limnologiques dominantes lors de la mise en place des dépôts ont une grande influence sur les communautés que l’on retrouve aujourd’hui dans le sédiment.
Au sein des deux principaux facies décrits ci-dessus, les métabolismes archées ont été analysés par le biais de séquençage d’ADN à haut rendement. Ces communautés semblent capables de gérer l’hypersalinité du milieu en utilisant les deux principales méthodes d’équilibrage osmotique, et peuvent aussi s’accommoder d’importantes concentrations en métaux lourds. Pour la première fois identifiée en Mer Morte, la méthanogénèse (à partir de composés méthylés) est un métabolisme clé du sédiment aragonitique. La fermentation de la matière organique résiduelle par des Halobacteria a lieu dans les deux types de sédiment. Des gènes de sulfato-réduction ont aussi été
vii détectés aux seins des métagénomes, en association avec la dégradation de biofilms microbiens et la présence de minéralisations de Fe et S.
A travers une comparaison avec un tapis microbien actif des bords de la Mer Morte, des clés de reconnaissances de minéralisations Fe-S d’origines microbiennes ont été acquises, et permettent de confirmer l’existence d’un cycle microbien du S en Mer Morte. La présence d’EPS témoigne de l’activité microbienne au sein du sédiment, et leur dégradation par sulfato-réduction produit des sulfures permettant la formation de minéraux précurseurs de la pyrite. Les minéralisations sont toutefois incomplètes, témoignant d’un processus incomplet de sulfato-reduction au sein du sédiment. De manière analogue, une relation intriquée entre EPS, nanoglobules et aiguilles d’aragonite suggère une influence microbienne sur la précipitation des carbonates de calcium en Mer Morte. Ces résultats ont de grandes implications en ce qui concerne la préservation de la matière organique et le développement des minéraux et de la porosité pendant les processus précoce de diagénèse en milieux hypersalins.
La mise en exergue d’une plus grande concentration d’organominéralisation de sulfure de fer dans un intervalle du début de l’Holocène, suggère une augmentation des taux de sulfato-réduction, pour cet intervalle précis. L’utilisation combinée de profiles élémentaires (sulfate) et isotopiques (C inorganique, S et O des sulfates) de l’eau interstitielle de la carotte, d’une description lithologique à haute résolution et des variations de signature lipidiques indiquent une période d’intense activité microbienne, en contexte relativement humide. Cet intervalle présente notamment un éventail de biomarqueurs indicatif d’une activité d’oxydation anaérobique du méthane. Entre autres, l’association de pentamethilycosene, archaeol et hydroxyarchaeol suggère la présence d’Archaea méthanotrophes du groupe ANME. L’oxydation anaérobique du méthane par ce groupe, associé avec des bactéries sulfato-reductrices semble avoir influencé certains proxies utiles aux reconstructions climatiques du début de l’Holocène.
En conclusion, la présence de vie et son extension sont aujourd’hui mieux contraintes dans les sédiments de la Mer Morte, et l’influence de l’environnement sur son développement est claire. La salinité est le principal vecteur de sélection des assemblages microbiens. Toutefois, l’apport d’eau douce en période humide permet le développement d’espèces aux métabolismes plus variés. Leur activité passée est aujourd’hui reflétée par les assemblages aujourd’hui présents dans le sédiment. En contexte lacustre, l’impact climatique sur les communautés microbiennes témoigne du besoin de régulièrement prendre en compte la microbiologie du sédiment, et également d’unifier les méthodes d’investigation au sein des projets ICDP. L’activité microbienne a un effet sur les signatures élémentaires et isotopiques de nombreux éléments, ainsi que sur la préservation de la matière organique, et ce même en conditions d’extrême salinité. Ce travail démontre donc la nécessité de systématiquement prendre en compte le rôle des microbes lors d’études paléoenvironnementales et lors de la construction de futurs modèles écologiques et climatiques.
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TABLE OF CONTENTS
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Remerciements ………………………………………………………………………………………….. i
Abstract …………………………………………………………………………………………………….. v
Résumé ……………………………………………………………………………………………………. vii
Table of contents …………………………………………………………………………………….... xi
List of tables ……………………………………………………………………………………..……… xv
List of figures …………………………………………………………………………………….……. xvi
I. Introduction: geomicrobiology and the Dead Sea Deep Drilling Project….. 1 1.1 Geomicrobiology and the importance of the microbial world to geology ………………………….. 3 1.2 The Dead Sea Deep Drilling Project ……………………………………………………………………………….. 7
II. Current microbial life in the Dead Sea sediment …………………………………. 17 2.1 Introduction ………………………………………………………………………………………………………………. 19 2.2 Impact of paleoclimate on the distribution of microbial communities in the subsurface sediment of the Dead Sea………………………………………………………. …………………………………………... 20 2.2.1 Introduction …………………………………………………………………………………………………………… 21 2.2.2 Materials and methods…………………………………………………………………………………………….. 22 2.2.3 Results ……………………………………………………………………………………………………………………..28 2.2.4 Discussion ………………………………………………………………………………………………………………. 32 2.2.4.1 Sediments as archives of environmental changes in the paleo Dead Sea …………….…... 33 2.2.4.2 Water column artifact or in situ communities ………………………………………………………. 35 2.2.4.3 Key parameters for the Dead Sea sediment subsurface ecology …………………………...... 36 2.2.4.4 Microbial community of halite-gypsum samples …………………………………………………… 39 2.2.4.5 Microbial community of aragonitic samples ………………………………………………………..... 40 2.2.4.6 Microbial communities in continental sediments ………………………………………………….. 42 2.2.5 Conclusion ………………………………………………………………………………………………………………. 43 2.3 Fluid inclusions as time capsules of chemistry and life ………………………………………………….. 44
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III. Metabolic potential of microbial communities in the subsurface of the Dead Sea ………………………………………………………………………………………………….. 59 3.1 Metagenomic studies in the Dead Sea subsurface…………………………………………………………. 61 3.2 Archaeal populations in two distinct sedimentary facies of the subsurface of the Dead Sea……………………………………………………………………………………………………………………………. 61 3.2.1 Introduction …………………………………………………………………………………………………………... 62 3.2.2 Methods ………………………………………………………………………………………………………………… 64 3.2.3 Results …………………………………………………………………………………………………………………... 66 3.2.4 Discussion ……………………………………………………………………………………………………………... 71 3.2.4.1 Metagenomics in poorly characterized environments …………………………………………. 71 3.2.4.2 Coping with harsh Dead Sea conditions ……………………………………………………………… 75 3.2.4.2.1 Salinity …………………………………………………………………………………………………………. 75 3.2.4.2.2 Toxicity of metals ………………………………………………………………………………………….. 76 3.2.4.3 Coping with deep sedimentary environments …………………………………………………….. 77 3.2.4.3.1 Fermentation ………………………………………………………………………………………………... 77 3.2.4.3.2 Methanogenesis …………………………………………………………………………………………….. 78 3.2.4.3.3 Sulfur reduction …………………………………………………………………………………………….. 79 3.2.4.3.4 Nitrogen cycle ……………………………………………………………………………………………….. 81 3.2.5 Conclusion …………………………………………………………………………………………………………….. 81
IV. Influence of microbes on the sedimentary record of the Dead Sea Basin………………………………………………………………………………………………………….97 4.1 Introduction …………………………………………………………………………………………………………….… 99 4.2 Geological setting ………………………………………………………..……………………………………………. 101 4.3 Material ……………………………………………………………………………………………………………………. 102 4.4 Methods ……………………………………………………………………………………………………………………. 105 4.5 Results ……………………………………………………………………………………………………………………… 105 4.5.1 X-ray fluorescence scanning ………………………………………………………………………………...... 105 4.5.2 Scanning electron microscopy ……………………………………………………………………………….. 107 4.6 Discussion …………………………………………………………………………………………………………….….. 111 4.6.1 Occurrence of traces of microbial activity in the Dead Sea sediment .……………………...... 111 4.6.2 Organic matter production and sulfate reduction .…………………………………………………… 115 4.6.3 Microbial effect of early diagenesis in the deep Dead Sea sediment…………………………... 118 4.7 Conclusion ………………………………………………………………………………………………………..……… . 122
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V. Influence of life in the Dead Sea geochemical record ………………………… 129 5.1 General influence of the biosphere on the carbon cycle of the Dead Sea (and its precursor lakes) …………………………………………………………………………………………………………….. 131 5.1.1 Introduction to the Dead Sea carbon cycle …………………………………………………………… 131 5.1.2 Carbon isotopes in the Dead Sea realm ………………………………………………………………… 132 13C signature in the core ……………………………………... 135
5.25.1.3 Anaerobic Aragonite oxidation precipitation of methane and itsdisturbed δ organic proxies in the Early Holocene Dead Sea ……………………………………………………………………………………………………………………….. 138 5.2.1 Introduction ………………………………………………………………………………………………………. 139 5.2.2 Geological setting ……………………………………………………………………………………………….. 140 5.2.3 Material ………………………………………………………………………………………………………………140 5.2.4 Methods …………………………………………………………………………………………………………….. 141 5.2.5 Results ………………………………………………………………………………………………………………. 143 5.2.6 Discussion …………………………………………………………………………………………………………. 147 5.2.6.1 Decoupling of isotopes with the precipitation-evaporation ratio in P6 ……………... 147 5.2.6.2 Information carried by biomarkers in the Dead Sea framework …..…………………… 149 5.2.6.3 Anaerobic oxidation of methane ……………………………………………………………………… 151
5.2.6.4 Diachronous 13CDIC and 34Ssulfate extrema ……………………………………………………….. 152
5.2.6.5 Tentative modelδ for enhancedδ microbial activity ……………………………………………... 153 5.2.7 Conclusion …………………………………………………………………………………………………………. 154
VI. Conclusion: the subsurface biosphere of the Dead Sea and lacustrine geomicrobiology …………………………………………….………………………………….….163 6.1 Towards the building of a unified model for geomicrobiology in the Dead Sea subsurface……………………………………………………………………………………………………………….…… 165 6.2 Reactions of microbial ecosystems to a changing environment: can the past of lakes be a key to the future …………………………………………………………………………………….…….. 168 6.2.1 From surface to bottom: microbial action in a changing environment ………….……... 168 6.2.2 When subsurface microbiology reflects climatic variations: geomicrobiology of lacustrine environments ………………………………………………………………………………………….. 170 6.2.3 Collaborative approach for understanding future changes in the aquatic biosphere of lakes and oceans ……………………………………………………………………………………… 171
Appendix ……………………………………………………………………………………………. 175
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LIST OF TABLES
Chapter II. Table 2.1 Sample description and principal geochemical characteristics ………………………………. 25 Table 2.2 Summary of primers for PCR amplification …………………………………………………………… 28 Table 2.3 Summary of PCR conditions …………………………………………………………………………………. 28 Table 2.4 Summary of number of clones picked and obtained OTUs ……………………………………… 32 Table 2.5 Sequences recovered from fluid inclusions …………………………………………………………….49
Chapter III. Table 3.1 Sequence information and diversity indexes for metagenomic samples …………………. 67 Table 3.2 M5RNA annotations for Archaea of sample AD ……………………………………………………… 68
Chapter V. Table 5.1 Principal chemical characteristics of lipid biomarker samples ………...…………………… 143
Appendix
Table A1 OTU definition and presence/absence in each sample ………………………………………….. 177
Table A2 OTU distribution and phylotypes definition …………………………………………………………. 180
Table A3 Calibration values for microthermometry analysis ………………………………………………. 180
Table A4 Temperatures of homogenization for 4 samples of the Holocene part of the core ...... 181
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LIST OF FIGURES
Chapter I. 1.1 Prokaryotic cell concentration and distribution in the sediment …………………………………… … 5 1.2 Idealized biogeochemical zone scheme in the marine sediment ……………………………………….. 6 1.3 ICDP drilled and planned lacustrine sites ………………………………………………………………………... 6 1.4 Location, elevation and geology of the Dead Sea Basin …………………………………………………….. 8 1.5 Unconformity between the Samra and Lisan Formations ………………………………………………… 9 1.6 Earthquake-related wavy structure within AAD laminations …………………………………………..10 1.7 Overview of DSDDP drilling sites ………………………………………………………………………………….. 11
Chapter II. 2.1 Overview of the sampling sites ……………………………………………………………………………………… 23 2.2 Detailed workflow for 16S rRNA gene sequence analysis ……………………………………………….. 27 2.3 DNA extraction samples and TOC and C/N profiles along the core ………………………………….. 29 2.4 CARD-FISH images of samples CT1 and CT3 ………………………………………………………………….. 31 2.5 Identified archaeal and bacterial OTUs ………………………………………………………………………….. 34 2.6 16S rRNA gene based phylogenetic tree of obtained Archaea sequences ……………………….... 37 2.7 Cluster analysis of identified phylotypes in the Dead Sea samples ………………………………….. 39 2.8 Halite rafts formation in the Dead Sea in December 2010 ………………………………………………. 45 2.9 Fluid inclusions and trapped organic matter in drilled halite ………………………………………….. 48 2.10 Putative Dunaliella cell enclosed in a fluid inclusion……………………………………………………… 48
Chapter III. 3.1 Archaeal classes from AD and GY …………………………………………………………………………………. 69 3.2 Heatmap of level 1 subsystems ……………………………………………………………………………………. 71 3.3 Heatmap of subsystems for osmotic adaptation and heavy metal tolerance …………………... 72 3.4 Heatmap of subsystems for methanogenesis, fermentation, nitrogen and sulfur metabolisms …………………………………………………………………………………………………………………… 74 3.5 Indications if active S cycle in the sediment core ………………………………………………………….. 77 3.6 SEM pictures of active organic matter and potential sulfate reduction ………………………….. 80
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Chapter IV.
4.1 Overview of sampling sites and material ……………………………………………………………………... 103 4.2 Aragonite needles, Dunaliella algae and EPS in the microbial mat………………………………….. 104 4.3 µ-XRF mapping of the microbial mat ……………………………………………………………………………. 107 4.4 Lithology and Fe-S content of the core …………………………………………………………………………. 108 4.5 Fe-S mineralization morphologies in the Dead Sea realm ……………………………………………… 110 4.6 EDX composition of Fe-S mineralizations ……………………………………………………………………... 112 4.7 Photographs of the cores exhibiting sulfur mineralizations…………………………………...... 114 4.8 SEM photographs of EPS morphologies in the core ……………………………………………………….. 117 4.9. Photographs of EPS structures in the mat and the core ………………………………………………… 119
Chapter V.
5.1 Carbon isotopes of DIC in the Dead Sea between 1963 and 1994 …………………………………….136 5.2 Carbon isotopes of aragonite in the core ………………………………………………………………………. 137 5.3 Location of the Dead Sea and the DSDDP drill sites ……………………………………………………….. 138 5.4 Lithological and geochemical profiles of the first 120 m of the core ……………………………….. 144 5.5 Lipid profiles of samples S17, S18, S19 and S21 ……………………………………………………………. 146 5.6 Proposed structures of mGD and macrocyclic archaeol……………………………..…………………… 147 5.7 Euhedral morphologies of Fe-S mineralizations in sample S17 ……………………………………... 148
Chapter VI.
6.1 Summary of the analysis carried out on the core core material within the PhD project…….168 6.2 Model for microbial assemblage settlement in the Dead Sea sediment …………………………... 169 6.3 Study of ecosystems evolution and reaction in a changing environment ……………………….. 172
Appendix
Fig. A1 Non-specific binding of tyramide on Dead Sea sediment …………………………………………. 182 Fig. A2 16S rRNA gene based phylogenetic tree of obtained Bacteria sequences …………………. 182 Fig. A3 Comparison of microbial diversity between clones amplicons and metagenomes……... 183 Fig. A4 Distribution of temperatures of homogenization from Holocene fluid inclusions ……… 183 Fig. A5 Halite block used for calibration of microthermometry …………………………………………… 183 Fig. A6 Carbon isotopes of AAD laminae of the microbial mat……………………………………………....184
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Fig. A7 Mass spectra of pentamethylicosene…………………………. …………………………………………... 184 Fig. A8 Partial chromatogram of fraction F3 for samples S17 to S21…………………………………….. 185 Fig. A9 Mass spectra of mGD and macrocyclic archaeol………..……………………………………………... 186 Fig. A10 SEM picture of greigite in P6 zone …………………………………………………….………………….. 187 Fig. A11 SEM picture of framboidal pyrite in P6 zone ………………………………………….……………… 187 Fig. A12 18 34S of pore water sulfate ……………………...……. 188
Linear relationship between δ O and δ
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Chapter I. Introduction: geomicrobiology and the Dead Sea Deep Drilling Project
“2+2 = 5” Hail to the Thief, Radiohead
1 Chapter I. Introduction
2 Chapter I. Introduction
1.1 Geomicrobiology and the importance of the microbial world to geology
The study of small life in the earth. The ancient Greek roots for the word “geomicrobiology” defines the purpose of this scientific domain. It encompasses the study of microscopic organisms, either prokaryotes or eukaryotes, in a geological realm. This geological realm may well be a mineral, a rock, soil, sediment, or any other geological object. The definition is actually wider, as it encompasses any microbial process influencing a geological object. Therefore, geomicrobiology deals with the degradation of organic matter and the synthesis of fossil fuels, with bioremediation in contaminated sites and bioleaching in ore deposits, with management of aquifers and drinking water quality. Obviously, it is a key scientific domain that has numerous applications in our modern society. It also covers purely theoretical scientific questions, in the domain of the origin, development and limits of life, on Earth and in the universe. The establishment of chemical or physical biologically derived signatures defines the search for the first traces of life on Earth. The understanding of the appearance of oxidized conditions on our planet, or the evolution of first eukaryotic cells requires knowledge of interactions of life with its immediate biological and mineral environment. The search for life on other planets demands the establishment of conditions and limits to its development, through for instance the study of extreme environments. The expansion of such science has however been restrained by the complexity of observing microscopic to nanoscopic features in the environments. In the last decades, advances in techniques of observation and DNA analysis, mostly guided by the field of medical sciences has allowed to establish geomicrobiology as a relatively common branch of geology, consolidating the already existing bridge between the living and the mineral worlds. There are very few environments where microbes are not present. A general rule has arisen from this: wherever there is liquid water, there is life. May the water be to the extremes of pH, temperature, radiation, pressure or activity. Archaea Pyrolobus fumarii can grow at 121°C (Blöchl et al., 1997). A barophilic bacterial strain close to the genus Moritella has optimum growth at 80 MPa (Kato et al., 1998). Deinococcus radiodurans can withstand up to 20 kGy of gamma radiation (Battista, 1997). The iron-oxidizing Archaea Ferroplasma acidarmanus grows at pH 0 (Edwards et al., 2000). Many archaeal species live at salt-saturation levels. As a result, the extensive colonization of media,
3 Chapter I. Introduction almost regardless of their physic-chemical characteristics, has shown that the potential for microbial influence on geology is everywhere. This is of major importance when it comes to the study of paleo-climates and paleo-environments. In most cases, such studies use the sedimentary record to tackle issues such as climate change, or mass extinction events. This is mostly done by combining different parameters influenced by the environment that are called “proxy”. These proxies may be elemental concentrations, microfossil appearances, isotopic ratios or physical properties of the rock. Some of these proxies are likely to be influenced by biological activity. Hence, they necessitate a very good control over their formation and preservation. Projects involving paleoclimatic and paleoenvironmental reconstructions thus need a good understanding of the microbiological processes at stake in the environment they are studying. One of the major sources for such studies is the ocean floor. It is also where the field of geomicrobiology has taken a major turn. Since 1986, the International Ocean Discovery Program (formerly Integrated Ocean Drilling Program) has been a major actor in the development of the field of geomicrobiology. Through the unique recovery of deep sediment, and well-adapted sampling and analysis methods, the notion of deep life has emerged, and with it, models for the distribution of microbes in the marine sediment (Fig. 1.1) and the suite of metabolic processes without light and oxygen (Fig. 1.2). Therefore, studies have allowed estimating that the sedimentary prokaryotic biosphere accounts for one tenth of the global biomass, and contributes almost equally as plants to the total fixed carbon (58 to 92 % of the plant-fixed carbon mass) making them key actors in the global carbon cycle (Whitman et al., 1998). Such studies have pushed the limits of life by for example emphasizing incredibly slow growth rates in the deep ocean sediment (Røy et al., 2012) and allowed to define new widespread phylogenetic groups (e.g. Miscellaneous Crenarchaeotic Group , Inagaki et al., 2003). The biomass has been tentatively quantified (Fig. 1.1A and B) and its tremendous influence on the global carbon cycle has been discussed (D’Hondt et al., 2004; Schippers et al., 2005; Kallmeyer et al., 2012). Among others, the role played by microbes in the formation and decomposition of methane hydrates (see AOM and methanogenesis in Fig. 1.2) is deeply investigated (e.g. Kvenvolden, 1995; Orphan et al., 2002; Inagaki et al., 2006; Knittel and Boetius, 2009) as several gigatons of methane carbon are estimated to be stored in continental margins (Milkov, 2004). These methane hydrates form
4 Chapter I. Introduction
Fig. 1.1: (A) Cell concentration along depth for different ocean sites and (B) geographic distribution model for cell number in the sedimentary column. Dots are calculated cell numbers at given sites. Figures taken from Kallmeyer et al. (2012). enormous sources of greenhouse gases in case of destabilization, and have already been evoked as the primary cause of mass extinction and global climate change in the past (e.g. during the Paleocene-Eocene Thermal Maximum (Dickens et al., 1995; Sluijs et al., 2007)). Thereafter, methane seep sites or gas-hydrate hosting shelves are prime target for the investigation of the ocean floor subsurface communities, and permit to link two main facets of IODP scientific objectives: the deep biosphere and environmental changes. Onto the continent, carbon cycles are less constrained. For example, Whitman (1998) estimated that the continental subsurface was 7 to 70 % as important as the ocean sediment one. The immense diversity of sites will possibly prevent any good prediction of microbial cell number, but the specificity of some of these environments have raised interest in the scientific community. Among them, lake sediments are prevailing target as they form excellent archive of past environments and are present in very varied size, geological settings and climate zones. Geomicrobiology of deep lacustrine sediment is only at its premises, and only very few lake sediments have been drilled with a planned microbiological investigation component in the ICDP framework (Fig. 1.3). However, the recognition of the importance of integrating such studies and a subsequent sampling protocol within the standard tools of scientific drilling has now emerged. This is highlighted by the new 2013 ICDP Science Plan, which encompasses for the first time subsurface microbiology as a main focus point of its program.
5 Chapter I. Introduction
Fig. 1.2 : Idealized biogeochemical zone scheme in the marine sediment modified from Konhauser (2007); *Anaerobic Oxidation of Methane (AOM) can work with other electron donor than sulfate reduced by sulfate reducers.
Lake El’gygytgyn
Lake Baikal
Bear Lake Great Salt Lake Lake Lake Van Ohrid Lake Qinghai Dead Sea Lake Lake Petén-Itzá Bosumtwi Lake Towuti
Lake Lake Titicaca Malawi
Laguna Potrok Aike Fig. 1.3: Map showing already drilled and planned ICDP sites with different levels of subsurface biosphere studies (larger dark gray and white dots, respectively). The smaller white dots indicate sites already drilled without special sampling for subsurface Figure 3 – Ariztegui, Thomas & Vuillemin biosphere studies. Notice that only ca. 30% of the so far studied sites have included geomicrobiological and subsurface biosphere sampling and subsequent investigations. From Ariztegui, Thomas & Vuillemin (2015).
6 Chapter I. Introduction
The first main results, mainly available for Laguna Potrok Aike (Vuillemin, 2013) and few from Lake Van (Glombitza et al., 2013) suggest important and varied communities, influenced not only by the geology and geochemistry of the sediment (as it is widely observed in the ocean sediment), but also by environmental changes and climatic variations, as lakes are more prone to these fluctuations than oceans. Peculiar responses of lakes subsurface biospheres seem inherent to the lake specificity, through diversities of location, response time to global changes, type and rates of sedimentation, and so on. As the turn towards continental subsurface microbiology is currently being taken, we intend to show through our research in the Dead Sea Basin, that even in environments hostile for life, it is relevant to undergo geomicrobiological study, and to integrate it into its geological history.
1.2 The Dead Sea Deep Drilling Project
Introduction to the geomicrobiology specific issues of the Dead Sea realm will be addressed in each of the following chapters. Here, I will rather give a broad view of the Dead Sea Basin geology and of the stakes and goals of a scientific drilling project in the Dead Sea. The Dead Sea is a terminal lake of the Levantine region (Fig. 1.4a). It now lies at the lowest point on Earth, at 427 below mean sea level (Fig. 1.4b). It is situated in the Dead Sea Basin, a pull-apart structure located within the Arava-Jordan Rift valley, between the Jordan Plateau and the Judean Mountains, and extending south towards the Red Sea in the Arava Valley, and North up to the Sea of Galilee (fig. 1.4c). In the past, difference in precipitation patterns and evaporation intensities have triggered extension and shrinking of the ancestor lakes of the Dead Sea (namely Lake Lisan, Lake Samra and Lake Amora, chronologically) within this basin. Such lake level variations have led to sediment deposition in areas that are today exposed (see Lisan Lake sediment key on Fig. 1.4c). These extensive variations have subsequently allowed the deposition and exposure of sediments. These easily reachable lake sediments constitute high resolution archives for paleoenvironmental, paleoseismic and paleoclimatic studies. Additionally, due to the nature of freshwater inputs, often occurring as flash floods, transport or organic debris is common and allows radiocarbon measurements. Moreover, uranium- rich aragonitic sediment dominating the Lisan and Samra Formation outcrops (Stein, 2001; Waldmann et al., 2007) extends dating capacities
7 Chapter I. Introduction
Fig. 1.4: (A) location of the Levantine region within the Mediterranean basin (NASA picture). (B) Elevation map of the Levantine region. Note that the Dead Sea Basin lies below sea level, and that the Dead Sea northern and southern basins were actually connected at the period of map design (1950’s). Red lines mark borders as defined by the UN Palestine Partition Plan for Palestine and the Armistice Demarcation Lines of 1949. Map taken from http://en.wikipedia.org/wiki/Geography_of_Israel (C) Geological map of the Dead Sea tributaries catchment area, taken from Neugebauer et al. (2014).
(Schramm et al., 2000; Torfstein et al., 2009) and permits good correlation with global climatic events, beyond two hundred thousand years. However, the halitic nature of the sediment dominating during interglacial periods prevents good preservation, as freshwater inputs readily dissolve the sediment. As a result, important hiatuses have been recorded in the outcropping record of the Dead Sea Basin (Stein, 2001 ; Fig. 1.5).
8 Chapter I. Introduction 275
Figure 3. The exposure of the Samra and Lisan Formations at Perazim Valley (west of Mt. Sedom). The Samra-Lisan depositional unconformity Fig. 1.5: Unconformity between the Samra Formation and the Lisan Formationappears between U-series ages of ~ 80 and 70 kyr. It is marked by sand and gravel. The upper part of the section, above the unconformity , with U- series ages of ca 80 kyr and 70 kyr, respectively.consist mainly of alternating laminae of aragonite and detritus, and hard benches Taken from Stein (2 that are made of gypsum. 001).
Lake Lisan water, and of clastic material transported by floods. The aragonite appears in thin (~ 0.5–1 mm thick) laminae Additionally, the Lake Lisan geographical existed between ~ 70 and and 15 kyr.tectonic During the location alternating with detrital of the laminae Dead of similar Sea thickness. makes it highly time of highest water level (> 180 m b.s.l.) the lake The detrital laminae are comprised mainly of quartz extended from the Sea of Galilee in the north to the gains probably of wind-blown origin, dolomite, calcite, susceptible to Hazeva seismic area in the activity. south (Figure 2). This The Lisan area Forma- is and indeed clay minerals known derived from to the be Cretaceous active wall- since ancient tion, which was deposited from Lake Lisan and its rocks. The gypsum appears in thin laminae, thicker lay- times. Particularly, surrounding biblical fan deltas consistsepisodes mainly of , chemicalsuch as ers (up the to 70 cm fall thick), of and the in disseminated Jericho form. Thick wall, have often aragonite and gypsum, which precipitated from the lake clastic layers are composed of sand, gravel and clay. tentatively been explained by scientists by earthquake events (Bentor, 1989; Ambraseys, 2006). Due to its high sedimentation rate and fine grained nature, the sediments have recorded this paleoseismic activity through wavy structures seismites notably observed in the highly laminated sediment of the Masada canyons (Fig. 1.6). The Dead Sea Deep Drilling Project (DSDDP) aims to obtain a continuous record of the Quaternary sediments of the Dead Sea Basin. This archive will help reconstructing past environments and climatic variation in a region at the crossroads of early hominid expansion, and that is considered as the religious and cultural cradle of society. In this way, the ICDP-sponsored DSDDP has drilled the lake floor at two key locations (Fig. 1.7a): the first one (5017-1) at 297,46 m in one of the deepest point of the lake (N31°30’28.98”, E 35°28’15.6”), and two other ones (5017-2 and 5017-3) on the shore of the lake, near the Ein Gedi spa (N 31°25’13.998”, E 35°23’38.4” and N 31°25’22.74”, E35°23’39.58” respectively). The drilling operations debuted in December 2010 and ended in March 2011, and obtained in total 722.65 m of sedimentary core,
9 Chapter I. Introduction
1m
Fig. 1.6: photograph of a seismite within the AAD facies of the Mishmar outcrop, near Masada. It is interpreted as the result of earthquake disturbance of unconsolidated laminae. with a recovery of 78.34 %. More details regarding recovery for each location can be found in Neugebauer et al. (2014). Such depth had never been attained in the Dead Sea sediment. The sedimentary archive obtained from the DSDDP is therefore unique and gives access to pristine material, highly valuable for geobiologists. Furthermore, the hypersalinity of its subsurface is for geomicrobiologists a unique feature as the Dead Sea constitutes an end member of hypersaline lakes on Earth, and as the study of subsurface communities in deep continental setting is very limited. Given the harshness of the lake water chemistry, and probably of its sedimentary pore water, the almost constant precipitation of evaporitic minerals, and the desire to use proxies derived from the drilled sediment for paleoclimatic reconstructions, several questions that will be discussed along this manuscript, arise. (1) Given the limited total number of cells in the present Dead Sea, and the extreme salinity of the water: is there life currently developing and/or surviving in the sediment? It is known that life exists in the surface of the sediments, thanks to freshwater springs that locally decrease salinity of the environment and allow microbial mats to develop (Ionescu et al., 2012; Häusler et al., 2014). Is this limited to the springs, do we see similar processes at depths, or is there another living community currently unknown?
10 Chapter I. Introduction
Fig. 1.7: Overview of drilling sites. (A) elevation map of the Dead Sea with names and location of ICDP drilled sites (taken from Neugebauer et al., 2014) (B) aerial photograph of the drilling platform on location 5017-1 (center of the lake). (C) photograph of the drilling platform at the coastal sites. Photo credit : ICDP
(2) If there is life, what are the sources for its development: how do communities manage to survive in those allegedly extreme salinities and low nutrient concentrations (phosphorus), and what kind of metabolism can be sustained in this hypersaline subsurface. Some thermodynamics limits have been set for the functioning of the main metabolic pathways at high salinities (Oren, 2010), are they validated in situ ? (3) Are potential microbes currently active in the sediment, or are they only dormant? If active, what kind of effect would they have on their environment and is it possible to find physical traces of their hypothetic activity? If traces of past microbial activity have existed, is it also possible to trace them, and would they have an influence on the sedimentary process ongoing in the subsurface. Among others, questions regarding precipitation/dissolution of mineral phases, and preservation or consumption of organic matter are at stake. Tackling these issues would allow constraining the carbon and sulfur cycles of the Dead Sea, and understand the impact microbial communities can have on the early diagenesis of hypersaline sediments. (4) Is microbial activity susceptible of disturbing proxies used for paleoenvironmental and paleoclimatic reconstructions? Sulfur isotopic ratios of gypsum from lake Lisan outcrops have been interpreted to bear signatures of bacterial sulfate reducing activity
11 Chapter I. Introduction
(Torfstein et al., 2005). Within the framework of the DSDDP project, it is highly relevant to set limits towards the use of some proxies potentially affected by life, albeit in settings hostile for life. If microbial disturbance exists, can one assess its trigger, its limits and its consequences? (5) Finally, wider questions bounded to the field of geomicrobiology could possibly be addressed: first, how do communities develop in the sediment? Do they originate from other environments, have they been transported from underground sources or do they relocate along the pore water of the sedimentary column? What is the actual control over their distribution? The precedent questions can be addressed within the specific framework of the Dead Sea subsurface, and possibly extended towards subsurface continental settings, when compared to the ocean realm, for example.
The following chapters will combine geological, microbiological, geochemical and biochemical datasets, in order to address these questions. Chapter II will focus on the currently living Dead Sea community and the information given by the community distribution, with 16S rRNA gene sequences as main tool. Chapter III will use metagenomic information to further understand the metabolic potential of the principal communities described in the sediment. In chapter IV, methods for the search for microbial traces will be discussed, as well as their meaning while the effect of past microbial activity will be questioned in chapter V, under the form of a multidisciplinary case study of a peculiar interval of the core.
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16 Chapter II. Current microbial life in the Dead Sea sediment
“Is there anybody out there?”
The Wall, Pink Floyd
17 Chapter II. Current microbial life in the Dead Sea sediment
18 Chapter II. Current microbial life in the Dead Sea sediment
2.1 Introduction
Determination of microbial presence using a simple microscope is complicated. Within sedimentary environments, this issue is even more complex as communities are often attached to particles and difficult to observe. Methods excluding visual identifications have thus been generally used in the sediments, and have been associated to fluorescent imaging to highlight the presence of microorganisms. Carried by pioneered work from Carl Woese (e.g. Woese, 1987), the identification of 16S rRNA gene sequences for determining microbial diversity is used routinely. Sanger sequencing, time and money consuming is currently being replaced by the higher-throughput sequencing methods, which allow not only the sequencing of targeted genes, but that of complete genomes present in a given environment. These techniques evolve very quickly and are constantly enhanced and tuned for better yields, time and money-wise. Reviews of these methods are available, and give a good view of pros and cons for the main ones (e.g. Thomas et al., 2012; Loman et al., 2012). In addition to the non-visual identification of DNA in a given samples, the need for understanding the spatial association of communities in a sample has arisen. With it, methods mostly developed from clinical investigations, have quickly established and are now used routinely. This is the case for 4',6-diamidino-2-phenylindole (DAPI) staining and fluorescent in-situ hybridization (FISH). They allow qualifying the presence of dead versus living organisms, and can target specific groups, either taxonomic or metabolic, using a sequence signature. FISH methods, which are based on the hybridization of a fluorescing probe to a targeted RNA sequence allows detecting RNA within a sample using a fluorescent microscope. With various probes, and combined with DAPI that binds to cellular material, the counting of microbial communities living in a given sample is made possible and gives more precise insight in a specific environment (e.g. Amann et al., 1995; Amann et al., 2001; Pernthaler et al., 2002; Pernthaler and Amann, 2004). These methods are routinely used in marine sediments and have become a corner stone of IODP investigations in microbial ecology (e.g. D’Hondt et al., 2004; Schippers et al., 2005; Inagaki et al., 2006; Biddle et al., 2008). It is with flow cytometry, a very efficient way of deciphering and counting microorganisms (McFeters et al., 1995; Porter et al., 1996). For such reasons, fixation of cells in order to allow the use of such dyes has been realized in the on-shore geobiology lab of the DSDDP, in Ein Gedi during the drilling campaign. Additional sampling for DNA extraction has been realized in the same lab. This material will be used as base material
19 Chapter II. Current microbial life in the Dead Sea sediment for the geomicrobiological investigation of the Dead Sea subsurface, addressed in the following part.
2.2 Impact of paleoclimate on the distribution of microbial communities in the subsurface sediment of the Dead Sea
A modified version of this chapter has been submitted to Geobiology as Thomas C., Ionescu D., Ariztegui D., and the DSDDP scientific party, Impact of paleoclimate on the distribution of microbial communities in the subsurface sediment of the Dead Sea
Abstract
A long sedimentary core has been recently retrieved from the Dead Sea Basin (DSB) within the framework of the ICDP sponsored Dead Sea Deep Drilling Project. Contrasting climatic intervals were evident by distinctive lithological facies such as laminated aragonitic muds and evaporites. A geomicrobiological investigation was conducted in representative sediments of this core. To identify the microbial assemblages present in the sediments and their evolution with changing depositional environments through time, the diversity of the 16S rRNA gene was analyzed in gypsum, aragonitic laminae and halite samples. The subsurface microbial community was largely dominated by the Euryarcheota phylum (Archaea). Within the latter, Halobacteriaceae members were ubiquitous, probably favored by their “high salt-in” osmotic adaptation which also makes them one of the rare inhabitants of the modern Dead Sea. Bacterial community members were scarce, emphasizing that the “low salt-in” strategy is less suitable in this environment. Substantial differences in assemblages are observed between aragonitic sediments and gypsum-halite ones, independently of the depth and salinity. The aragonite sample, deposited during humid periods when the lake was stratified, consists mostly of the archaeal MSBL1 and bacterial KB1 Candidate Divisions. This consortium probably relies on compatible solutes supplied from the lake by halotolerant species present in these more favorable periods. In contrast, members of the Halobacteriaceae were the sole habitants of the gypsum-halite sediments which result from a holomictic lake. Although the biomass is low, these variations in the observed subsurface microbial populations appear to be controlled by biological conditions in the water column at the time of sedimentation, and subsequently by the presence or absence of stratification and dilution in the lake. Since the latter are controlled by climatic changes, our data suggests
20 Chapter II. Current microbial life in the Dead Sea sediment a relationship between local lacustrine subsurface microbial assemblages and large- scale climatic variations over the Dead Sea Basin.
2.2.1 Introduction
The present Dead Sea is located in the Dead Sea Basin at the border between Israel, Jordan and the Palestinian Authority, (Fig. 1a). Its sediment record spans several hundred thousand years of climatic variations (Torfstein et al., 2009), making it a prime target for geological investigations. To better understand the climatic response of this currently water stressed region, over the last glacial-interglacial cycles (Goldstein et al., 2011; Neugebauer et al., 2014), the ICDP sponsored Dead Sea Deep Drilling Project (DSDDP) retrieved the longest sedimentary core to date from the deepest point in the lake. Since more than 50 years now, the Dead Sea water budget has been largely negative, resulting in a massive drop of the shoreline at an increasing rate which in the last decade exceeded one meter per year. Since 1979 the lake has been fully mixed and oxic with the exception of short periods of stratification. Halite is almost constantly precipitating and the total dissolved salt concentration has reached one of the highest levels on Earth with up to 348 g. L-1 (Oren and Gunde-Cimerman, 2012). Interestingly, the Dead Sea chemistry is characterized by a high concentration of divalent cations (~2 M Mg2+ and ~0.5 M Ca2+), which make it a specifically harsh environment even for microorganisms (Oren, 2001). Indeed, except for some oases of life located at the outflow of groundwater springs, the microbial cell number is extremely low in the present Dead Sea water (<104; Ionescu et al., 2012). The only eukaryote and primary producer in the Dead Sea water column is the green alga Dunaliella, which blooms periodically when the surface layer has been diluted enough by heavy rainfalls over the Jordan catchment area and the surroundings of the Dead Sea (Oren et al., 1995). The extreme salinity of the modern Dead Sea mainly allows for the existence of halophilic Archaea of the Halobacteriaceae family (Oren, 1994; Bodaker et al., 2010). This dominance is probably due to the utilization of the low energetic cost “high salt-in” osmoregulation strategy in which cells balance the high external salinity by pumping in K+ ions. In contrast, the “low salt-in” strategy which is more common in halotolerant prokaryotes and halophilic eukaryotes relies on the synthesis or uptake of organic molecules (called compatible solutes) that act as osmoprotectants. Being more energy demanding, this strategy is less suited for
21 Chapter II. Current microbial life in the Dead Sea sediment extreme environments like the Dead Sea, as it leaves little or no energy available for growth (Oren, 2011). The Dead Sea was not always as harsh for life as it currently is. Humid periods allowed for the establishment of meromictic episodes providing a sufficiently diluted epilimnion and enough nutrients for life to develop, as shown by remnants of diatoms in its sediment (Begin et al., 1974; Thomas et al., 2014). Sedimentary and geochemical data from outcropping sediments suggest that the hypolimnion may also have hosted anaerobic bacterial communities able to metabolize and take part in the complete carbon and sulfur cycle of the paleo Dead Sea (Luz et al., 1997; Torfstein et al., 2005). Recently, the analysis of archaeal metagenomes from deep aragonitic and gypsum samples of the DSDDP core has revealed the occurrence of different communities in each facies, surviving using different metabolic pathways (Thomas et al., 2014). Nevertheless, low read coverage in these metagenomes resulted in the retrieval of only few 16S rRNA gene sequences thus preventing full description of the microbial assemblage. To understand the extension of life development to the hypersaline subsurface of the Dead Sea, and the mechanisms leading to its variability, selected intervals representing specific periods of lake sedimentation have been sampled in a unique, 456 m-long, core. This record was retrieved during the DSDDP campaign in winter 2010-2011 (Fig. 1b-c) at more than 300 m below the lake surface. Pore waters of these sediments trapped the chemistry of the paleo-lake waters, reflecting the actual conditions for life development, as discussed by Lazar et al. (2014). We compare the microbial communities from the core material with those of a microbial mat from the shore of the present Dead Sea. This mat serves as a reference of an active sedimentary microbial assemblage under the current chemistry of the basin. We aim to provide a comprehensive view of the Dead Sea subsurface biosphere, its putative limits and the potential influence on the biomass of the various sedimentary facies.
2.2.2 Material and methods
Sampling
Except for sample CT2 (Table 1), core-catcher sediments have been sampled onshore directly after drilling (December 2010). Specific care was taken to avoid any external or trans-contamination as detailed later-on. Right after each drilling shift, cores and core-
22 Chapter II. Current microbial life in the Dead Sea sediment
Fig. 2.1 (a) and (b) Landsat images of the Dead Sea in the Levantine area and zoom up of the Dead Sea and sampling locations. Yellow star indicates the drilling site where the core presented here has been retrieved. Red star shows sampling location of the microbial mat, in En Qedem; (c) Photograph of the ICDP drilling barge on site; and (d) Dead Sea shore pool where the microbial mat has been sampled. Red colour is due to numerous Dunaliella pigments in the water and on the microbial mats covering the ground. Pool is 15 m long. catchers were brought to the shore and stored in a cooling room (4 °C). Core-catchers were then opened in a specially arranged geomicrobiology lab and sampled for microbiological studies using pre-cut and autoclaved syringes. Mini-cores were then put into sterile tubes and stored in a freezer at -8 °C. Sample CT2 was sampled during core opening, in June 2011 at the GFZ in Potsdam. The core from which it originates was stored in a cooling room at 4 °C and sampled for geomicrobiological investigation immediately after sawing. To prevent any contamination from circulating fluids within the core liner or from the saw, the first centimeter of a 5 centimetre-long core was removed and the rest (inner part) was rapidly collected. All samples were stored at -8 °C before further processing.
23 Chapter II. Current microbial life in the Dead Sea sediment
The samples from the microbial mat were collected below a salt crust in a temporal pool along the western Dead Sea shore, near En Qedem (Fig. 1d) by cutting a cubical section of the sediment. The sediment was then wrapped in Parafilm® and stored in the freezer before further processing. Subsampling was done in a clean lab using sterile tools. Samples were subsequently separated into a red layer (M1) and the underlying white one (M2). Aliquots from the geomicrobiological samples were used for total organic carbon and N measurements, as well as for lithological description, using sterilized material. Porewaters were analysed from core-catchers and were collected immediately after geomicrobiological sampling. Table 1 summarizes the main features of the different samples.
Organic C and N determinations
Samples for TOC and N were multiply rinsed with ultrapure water, centrifuged and oven-dried at 40 °C. After acidification, they were measured on a PE 2400 Series II CHNS/O Elemental Analyser at the University of Geneva, with a reproducibility of 0.11 % and 0.01 % error margin for C and N respectively.
Catalyzed Reporter Deposition – Fluorescence In Situ Hybridization (CARD-FISH) Thirty-one samples from the first 100 m were fixed in the field lab using 3 % formaldehyde with PBS and then rinsed and stored in ethanol as previously described (Vuillemin et al., 2013). Subsequently, 300 µL from each sample were vortexed (30 s) and sonicated (30 s) three times to detach cells from the fine sediment particles. The cell-containing supernatant was added to 25 mL of sterile PBS, which was filtered on polycarbonate filter (0.2 µm pores size 47 mm Ø) and then rinsed with milliQ water. Hybridization was performed according to Ishii et al. (2004) using probes EUB338 for Bacteria (Amann et al., 1990) and ARCH915 for Archaea (Stahl and Amann, 1991). The NON338 probe (Amann et al., 1995) was used to test for non-specific hybridization. Photographs were taken using a Zeiss Axioplan microscope with a 100 W Hg lamp and filters for DAPI and Alexa 488. To test for non-specific binding of the tyramide dye to the sedimentary material, the protocol was also carried out on several samples without adding any probe.
24 Chapter II. Current microbial life in the Dead Sea sediment
Depth Sample Deposition Salinity Na+ Ca2+ Mg2+ Cl- SO 2- Sample Lithology 4 (m) Type environment (%) (mM) (mM) (mM) (mM) (mM)
current core CT1 0.24 Halite holomictic Dead 33.60% 1630 392 1825 6320 5.8 catcher Sea
alternating Meromictic CT2 2.74 core aragonite and mud Holocene Dead 33.21% 1726 429 1745 6177 4.2 laminae Sea
End of the core Pleistocene CT3 90.64 gypsum 24.90% 1795 74 321 5438 23 catcher holomictic Lake ~ Lisan
Pleistocene core CT4 206.53 Halite holomictic Lake 33.86% 1555 442 1827 6389 4.5 catcher ~ Samra
Current red mini gypsum and halite microbial mat at M1 0.01 36.90% 2120 583 1586 6110 18.9 core in EPS the Dead Sea shore
Aragonite mini laminae right M2 0.07 aragonite ------core below the microbial mat
Table 2.1 Sample description and principal geochemical characteristics. Data from sample CT2 have been retrieved from Nissenbaum (1975) as expected to be similar to the deep water mass before the 1979 turnover evidenced by the beginning of halite precipitation in the top meters of the core. M2 concentrations were not measured but are considered similar to M1 since it is only 2 mm below.
DNA extraction and sequencing
DNA extraction is a critical step due to the low DNA content and the peculiar chemistry of the Dead Sea sediments. Therefore, we provide here the complete workflow used to assure the reliability of the retrieved sequences (Fig. S1 supp. mat).. Selection of the best-suited DNA extraction protocol was done after various trials, and subsequently tuned. Before discarding a DNA extraction protocol, combination of results from Nanodrop® measurements (both DNA quantity and quality ratios), as well as from nested PCR were examined. Each protocol was tested on multiple samples to avoid biases inferred by the specific facies or chemistry of the sample. Once the protocol was selected, a sample spiked with control-DNA was extracted in parallel to test for the coextraction of inhibitors for downstream processes. Samples for which replicates could
25 Chapter II. Current microbial life in the Dead Sea sediment be produced from the extraction to the sequencing step were kept and their results are presented in the following sections. Other samples are considered as void of analyzable DNA. A second PCR was systematically needed before obtaining agarose-gel positive bands and clonable products. All positive clones were picked and sequenced. The chosen DNA extraction, protocol was based on a phenol–chloroform extraction method modified from Ionescu et al. (2009), and proven to work in the challenging Dead Sea sediments (Ionescu et al., 2012). Cells were extracted from 0.5 g of sediment after multiple cycles of PBS rinsing, 30 sec sonication and quick centrifugation. Subsequently the samples were then incubated for 20 min at 95 °C in 0.5 mL lysis buffer (0.1 M Tris, 50 mM EDTA, 100 mM NaCl and 1 % SDS pH 8). 500 µL of Phenol: chloroform: isoamyalcohol was added and the samples centrifuged at 16 873 x g after incubation for 10 min at room temperature. Supernatant was extracted again with the same process and the upper phase collected using Phase Lock GelTM (PLG) tubes (5 Prime). The aqueous phase was further cleaned twice with 0.5 mL 24 : 1 (v : v) chloroform : isoamylalcohol after which the DNA was precipitated overnight at -20 °C in 1 volume of isopropanol and 2 % (final volume) of 3 M sodium acetate (pH 5.5). After a 30 min centrifugation at 16 873 x g Pellets were washed with 0.5 mL of 70 % ice cold ethanol, dried and finally dissolved in 10 µL molecular grade water. The QIAEX II Gel extraction Kit (Qiagen) was used to remove salt and purify DNA fragments according to the manufacturer instructions. DNA was quantified using a Nanodrop® ND-1000 Spectrophotometer (Witec AG). To recover sufficient amount of DNA for cloning, an initial PCR done using the universal primers 4F and Univ1492R (Dong et al., 2006) was followed by a nested PCR with 1 µL of PCR using the 3F & 9R primers for Archaea (Jurgens et al., 1997). For Bacteria, we used 27F and 1492R followed by 341F and 907R (Lane, 1991; Weisburg et al., 1991; Muyzer et al., 1993) (Table 2). DNA fragments were amplified using the protocol presented in Table 3. After purification with High Pure® PCR product Purification Kit (Roche Diagnostic SA), the products were cloned using the TOPO TA Cloning Kit (InvitrogenTM by Life TechnologiesTM) following the manufacturer’s instructions. Positive clones were verified by PCR and subsequently prepared for sequencing using the BigDye® Terminator v3.1 Cycle Sequencing Kit (Applied BioSystems) with primers D4 and R5. Sequencing was performed on an ABIPRISM® 3130xl Genetic Analyzer (Applied BioSystems, Hitachi).
26 Chapter II. Current microbial life in the Dead Sea sediment
16S rRNA gene sequence analysis
Sequences were cleaned, assembled, trimmed to around 550 nucleotides and aligned using CodonCode Aligner© v.3.7.1 (CodonCode Corporation) and Seaview® v.4.0.0 (Gouy et al., 2010). Sequences were checked for chimeras using Bellerophon (Huber et al., 2004) and were subsequently identified using the megx® Geographic-BLAST (Lombardot et al., 2006) and the SILVA rRNA database project (Quast et al., 2013). Operational Taxonomic Units (OTUs) at a 3 % cut-off (97 % similarity) were defined using mothur (Schloss et al., 2009 ; supp. material Table S1). Reference sequences were downloaded from the SILVA© database and the NCBI genbank®. All the sequences (samples and reference) were uploaded to the Ribosomal Database Project page (Cole et al., 2009) and a phylogenetic tree was built through the Weighted Neighbor Joining method (Bruno et al., 2000) for representative archaeal and bacterial phylotypes. All sequences have been deposited to the BankIt1757557 library under references KM587723 to KM587887.
Fig. 2.2 Detailed workflow used for 16S rRNA gene sequence analysis. Green coloured arrows correspond to flow in case of succesful results while red ones correspond to steps in case of negative ones.
27 Chapter II. Current microbial life in the Dead Sea sediment
Table 2.2 Summary of primers used for PCR amplification of 16S rRNA gene for Bacteria and Archaea
2.2.3 Results
General description of the core and samples
The studied core can be divided into four main sections based on the dominant lithologies (Fig. 2). From 457 m (deepest point) to around 320 m, the core is mainly composed of alternating laminae of aragonite and detrital mud (AAD facies). Occasionally, laminae can reach a few centimetres in thickness. They can be intercalated by intervals of detrital mud, often fining upwards. From 320 to 200m, the core is mainly composed of halite either as rock salt, or mixed in detrital mud with occasional intercalations of AAD intervals. From 200 to 100, AAD dominates again, and is rather monotonous. An interval dominated by pure gypsum is found from 100 m to 90 m just before halite deposits resume again and remain dominant to the top of the core, often mixed with marly detritus. A more complete and detailed lithological description can be found in
Table 2.3 Summary of PCR conditions for each Neugebauer et al. (2014). primer set.
28 Chapter II. Current microbial life in the Dead Sea sediment
The modern microbial mat from the shores of the Dead Sea is composed of gypsum and halite crystals mixed with some detritic clay material embedded in a microbial mucilage matrix of External Polysaccharide Substances (M1; Fig. 2E). Sample M2, lying immediately under M1 (Fig. 2E) is composed only of aragonite needles as also observed in the aragonite laminae of the AAD facies in the core (Fig. 2B).
TOC and C/N ratio
Total organic carbon varies between 0.02 % and 2.46 % (Fig. 2) averaging 0.75 % (of total dry weight). Due to the large sampling interval it is difficult to identify consistent trends in the data, especially below 200 m. However, a general increase of TOC from 0.5 % to more than 2 % appears between 200 m and 100 m depth. From 100 m to the
Fig. 2.3 Left panel (a) shows lithological description of the studied core modified from Neugebauer et al., (2014) and TOC and C/N profiles. Many attemps of DNA extractions at different depths failed. Their stratigraphical position along with their lithology are indicated with red crosses. Analogously, successful ones are indicated with green ticks. Letters correspond to pictures in panel b. Right panel (b) displays pictures of samples from which microbiological samples have been taken: (A) Surficial Halite CT1; (B) AAD facies at 2.74 m CT2; (C) Gypsum at 90.64 m CT3; (D) Halite at 206.53 m CT4; and (E) Microbial mat sample. M1 was taken in the surficial red laminae, while M2 was sampled immediately below CT8, in the white aragonitic layer. Scale bar always indicates 0.5 cm.
29 Chapter II. Current microbial life in the Dead Sea sediment surficial sediments the values drop, stabilizing between 0.5 and 1 %. The C/N ratio at depths over 200 m also shows fluctuating values which peak between 200 m and 67 m. Up to the surface, the C/N ratio reached minima values although some variations can be observed at 23 m. Overall, the C/N ratio extends from minimum values at 65 m (3.50) and a maximum at 109 m of 23.67.
CARD-FISH imaging
The NON338 negative control probe gave positive results for 29 out of 31 samples and similar false-positive results were obtained from the sample incubation with the tyramide dye. The shallowest halite sample, and the gypsum sample were the only ones which did not produce a false positive result (Fig. 3). Therefore, we present only images for CT1 and CT3, which serve to highlight the presence of microbes within the sediments of the Dead Sea, down to at least 90 m below lake floor (mblf). Given these methodological limitations, the CARD FISH images were not used for cell enumeration. Archaeal and bacterial cells were identified in the shallow halite sample, with relatively low density, pertaining to the nature of the sediment (halite crystals). For the gypsum sample, the abundance of sedimentary particles did not prevent cell visualization. Archaea were more abundant than Bacteria and the total microbial population was denser than in the halite sample.
Community analysis
From the six sedimentary samples investigated (4 samples in the core and two in the microbial mat), 188 OTUs were obtained out of 624 clones (Table 4; Table S2 of supp. material). All sequences of Archaea cluster within the Euryarcheota phylum and are almost all related to organisms from hypersaline environments, varying from moderate to extreme halophiles. The most represented phylotype is of the Halorabdus genus, with 82 OTUs. The second most abundant phylotype consists of uncultured Halobacteriaceae (9 OTUs). Other identified genera are Halonotius (4 OTUs), Halomicrobium (3 OTUs), Haloplanus, Halobacterium and Haloterrigena (1 OTU each). Other than Halobacteria, two additional archaeal classes were identified within the Dead Sea sediments sequences: Thermoplasmata MSBL1 Candidate Division (23 OTUs) and Methanomicrobia (1 OTU).
30 Chapter II. Current microbial life in the Dead Sea sediment
The microbial communities from the halite-dominated sediments sampled from the shoreline microbial mat and from the core show strong similarities (Fig. 4). Both contain sequences linked to uncultured Halorhabdus generally identified in the Dead Sea water, as well as in other hypersaline water and sedimentary environments. Some sequences belong to an uncultured Halobacteriaceae phylotype (Fig. 5), closely related to sequences retrieved from the hypersaline sediments of Lake Kasin in southern Russia (Emmerich et al., 2012). Finally, the largest number of archaeal OTUs from the halite- gypsum sediments cluster in a poorly defined group (Dead Sea sediment group; Fig. 5), tentatively related to uncultured Archaea close to the Halorhabdus genera. Closest blast sequences have been retrieved from the hypersaline lake Tuz in Turkey (Mutlu et al., 2008), and from hypersaline alkaline salterns in East African lakes (Grant et al., 1999). The sequences retrieved from the AAD sample are unique, strongly separating this sample from the others. It is largely dominated by sequences associated to the MSBL1 candidate division sequences (92%). Sequences similar to uncultured Methanomicrobium and Halobacteriaceae from a Tunisian solar saltern complete the assemblage. Finally, the archaeal population in the microbial mat is rather homogenous
Fig. 2.4 CARD-FISH images of samples CT1 (A-C) and CT3 (D-F) showing the negative NON338 probe response (A and D), the ARCH915 hybridized cells in green corresponding to Archaea (B and E) and the EUB338 hybridized cells in green corresponding to Bacteria (C and F). In yellow are particles and in blue are remnant DAPI-stained cells (living Archaea/Bacteria or dead cells)
31 Chapter II. Current microbial life in the Dead Sea sediment
and similar to the halite and gypsum samples of the core sediments; with a great dominance of Halobacteriaceae related clones among which Halorhabdus are the most represented sequences. Surprisingly, a low richness is found in the mat, along with few changes between the red halite-gypsum crust and the aragonite lamina. PCR products for bacterial DNA clustered into 46 OTUs grouped in 9 bacterial classes. Sample CT2 has the highest richness in the core, displaying 15 OTUs, with 3 of them related to the Deferribacterales order and 12 affiliated to the Candidate Division KB1. This Candidate Division is also largely represented in sample M2 of the mat (14/23 OTUs). The aragonitic mat sample is overall the richest sample with 22 OTUs across five known phylotypes. Sequences related to the Halanaerobiaceae family make almost a third of its bacterial 16S rRNA gene library. Only 3 bacterial OTUs could be retrieved from the surficial halite sample. They are associated to halophilic or halotolerant members of the Deinococci, Nitrospira and δ-proteobacteria classes (Fig. 4B and Fig. S1). At depth, gypsum and halite did not result in any bacterial amplicons although few cells are visible in the CARD-FISH images (Fig. 3). Only three bacterial OTUs, from Bacteroides (2 OTUs) and Rhodospirillaceae Limimonas were obtained from the mat crust sample M1. Similarity to known archaeal sequences varies a lot, with identity percentages ranging from 97.8 % for Halorhabdus related sequences, to less than 71 % for Halobacteriales related to a Halomicrobium found in the shallow halite sample. Similarly diverse identity percentages are observed for bacterial sequences (from 70 % to 99 % for KB1 Candidate Division associated sequences).
Table 2.4 Summary of number of clones picked and obtained OTUs for each sample
32 Chapter II. Current microbial life in the Dead Sea sediment
2.2.4. Discussion
We present here the first evidence of a microbial community in the deep subsurface of the Dead Sea. In the following sections, we will describe and discuss the dataset with respect to the interactions of these communities with their specific environment. To do so, we will first address the unique characteristics of the Dead Sea, and later the key features of the various microbial assemblages. Last, we will emphasize its relevancy in the framework of our sedimentary record.
2.2.4.1 Sediments as archives of environmental changes in the paleo Dead Sea
The various lithological facies described in the core are representative of the prevailing limnological regime at times of deposition (Neugebauer et al., 2014). Halite deposits are the result of intense evaporation in holomictic conditions. They mark the harshest conditions in the Dead Sea water column. These deposits dominate during the Holocene (Marine Isotopic Stage 1 - MIS1) and MIS5 interglacial periods. Gypsum precipitates as thick layers or disseminated laminae. This occurs during increasingly arid conditions, probably following major changes in the limnological regime from stratified to mixed conditions (Torfstein et al., 2008). Finally, the AAD facies is interpreted as a rhythmic deposit (Fig. 2b) occurring principally during glacial periods, when the lake is stratified (Stein et al., 1997). Freshwater flows to the lake bringing detrital material that is rapidly deposited. The subsequent precipitation of the aragonitic laminae is still under discussion but appears to be linked to the input of carbonate rich freshwater from the Jordan river and ephemeral wadis, that reacts with the highly concentrated Ca2+ ions from the lower Ca-Cl brine when evaporation or phototrophic activity increases alkalinity (Kolodny et al., 2005). The climatic and evaporation/precipitation ratio changes are also outlined by the profiles of the TOC and C/N ratio, with an additional insight on the fate of organic matter. While organic matter accumulates in the sediment during aragonite precipitation periods, the C/N ratio indicates that this organic matter is mainly allochthonous compared to the one measured in the upper part of the sedimentary core. Values lower than 10 are typical of autochthonous organic matter, most often derived from phytoplankton while vascular plants display C/N ratios above 20 (Meyers, 1994). In recent aquatic sediments, early diagenesis can potentially increase the C:N ratio with depth (Giani et al., 2010) through preferential removal of N from
33 Chapter II. Current microbial life in the Dead Sea sediment
Fig. 2.5 Identified archaeal (A) and bacterial (B) phylotypes using 16S rRNA gene sequences and defined OTUs using a 97% cutoff. Numbers represent the amount of OTUs identified for a given phylotype. This identification is based on environmental blast. labile organic matter. Here, no major deviation is observed with depth. Instead, an increasing amount of organic matter, with more contribution of land plants, can be seen in the Dead Sea sediment during humid periods when AAD dominates. In contrast, during periods of halite precipitation, little impact of allochthonous organic matter can be seen, indicating arid conditions in the catchment. As their sedimentary counterpart, organic facies are thus representing specific climatic conditions, and based on the
34 Chapter II. Current microbial life in the Dead Sea sediment current chronology these sections can be preliminary correlated to the last two glacial- interglacial cycles (Neugebauer et al., 2014). Considering the strong compaction of the samples, the very high sedimentation rate, and the fact that almost all the halite intervals are interbedded with relatively thick, dense marly layers, we speculate that the general paleo Dead Sea chemistry was preserved in the sediment pore waters. Diffusion, although acknowledged to be present is limited by salt layers that act as seals between marly intervals (Lazar et al., 2014). Subsequently, the microbes investigated in each layer should have remained in environments relatively similar to those experienced at the moment of deposition, except for their own disturbance derived from potential metabolic activity. The latter has strong implications on the geomicrobiological interpretations that can be drawn for this environment.
2.2.4.2 Contamination, water column artifact or in situ communities?
The use of core catchers for microbial sampling, and the potential for infiltration of drilling material or lake water in the most permeable samples, such as those containing halite (Fig. 2A and D) may raise some questions. Extreme care was taken during sampling procedure, notably by making sure that none of the sediment samples was ever in contact with parts of the coring devices such as the liner or the core catcher diaphragm. Our results support the absence of contamination. Drilling fluid consisted of Dead Sea water and sterile industrial guar gum. Its contaminants may thus be members of the Dead Sea water column, such as Halobacteria. However, the workflow we used was designed to allow us to rule out the option of contamination. Hence, it secures the autochthonous origin of the retrieved communities. If contamination pertaining to the drilling material had occurred, there would be no reason for a differential pattern of DNA retrieval along the core as the microorganisms in these fluids would outnumber those in the subsurface environment of the Dead Sea. Furthermore, samples spiked with known DNA were used in parallel for all samples. All sequences of the known DNA were obtained from all spiked samples regardless of the sample type, allowing us to rule out inhibition inherent to the sediment physics and chemistry. Thus, the fact that no DNA could be obtained from a good number of samples (Fig. 2) does infer that contamination is not the source for organisms obtained in the cored sediments. Additionally, this shows that the biomass in the sediment of the Dead Sea is very low.. Conclusively, we argue
35 Chapter II. Current microbial life in the Dead Sea sediment that contamination from the water column, drilling fluids or circulating fluids can be ruled out. Other sources of external deviation from the original sedimentary community may arise from PCR bias, especially given that a nested PCR was needed for obtaining sufficient DNA. Protocols were thoroughly reproduced in order to minimize such issues. Sample source ( either from the core catchers, the core or the mat) should not lead to change in microbial community profile as the downstream processing of all samples was similar.. Before validation, positive and negative results were reproduced to further discard possible cross-contamination.
2.2.4.3 Key parameters for the Dead Sea sediment subsurface ecology
Previous studies on the Dead Sea microbiology focused mainly on the water column (Nissenbaum, 1975; Oren, 1983b; Bodaker et al., 2010; Rhodes et al., 2012) or the very shallow sediments (Ionescu et al., 2012). Recent investigation of archaeal metagenomes has shown the potential for retrieval of DNA in the deep Dead Sea sediment (Thomas et al., 2014). The activity of the microbial community in the sediments is however still hard to quantify. Monitoring of microbial activity was planned through the detection of ATP production in the sediment (Vuillemin et al., 2010); however, results remained negative even when used on the microbial mat samples M1 and M2, which can hardly be inactive given the observed EPS production and eukaryotic cell presence (not shown here). Research on luciferin-luciferase systems has highlighted the tendency for inorganic salts to inhibit luminescence at high anionic concentrations (Gilles et al., 1976; Rodionova and Petushkov, 2006). We thus hypothesize an inhibition of the enzyme under the Dead Sea salinity, making luciferase-dependent ATP-tests poorly suited for such projects. Direct counting techniques using fluorescence in situ hybridization turned out to be inefficient in most of the Dead Sea sediments, mainly due to the strong background fluorescence created by the unique and highly concentrated composition of the pore water, and the very fine-grained lithologies (Fig. S2). This was still the case after repeated cleaning of the sediment. Fortunately, CARD-FISH images could be acquired for two of the 5 samples presented in this study. Negative controls testify to the validity of these pictures and attest that cells are present at least to a depth of 90 m in the core. Whether these cells are alive or active is still a matter of debate.
36 Chapter II. Current microbial life in the Dead Sea sediment
Fig. 2.6 16S rRNA gene based phylogenetic tree of the Archaea representative sequences for phylotypes defined in this study. Tree was built using the Weighted Neighbor Joining method. Scale 10% of estimated difference in nucleotide sequences. Bars on the right panel identify main archaeal classes and major clusters identifiable in the Dead Sea sediments
37 Chapter II. Current microbial life in the Dead Sea sediment
Euryarcheota is the only represented archaeal phylum in the sediment. The archaeal population in the Dead Sea sediment is largely composed of obligate halophiles, what would suggest that the extreme salinity of the Dead Sea sediment is the main factor controlling archaeal ecology. However, samples CT2 and M2 can be distinguished from the halite-gypsum samples (Fig. 6). The dominance of the bacterial KB1 Candidate Division, and the low abundance of Halorhabdus related sequences, marks this difference. The lowest measured salinity was obtained from the gypsum sample while the values measured in CT2 and M2 are similar to those measured in the shallow halite sample. Hence, salinity is not the sole determinant of microbial distribution. Similarly, one could expect depth or deposition time to be of prime influence, but the relative similarity between the microbial communities of the shallow CT1 and M1, and deep CT3 and CT4 samples goes against such inference. In the future a higher sampling resolution with depth, coupled with improved DNA extraction and amplification techniques as well as activity determination protocols that overcome the Dead Sea chemistry may provide a better understanding of the main factors controlling the microbial ecology of the Dead Sea sediment. As a first approach, however, we compared and found a good match (Fig. S3) between the 16S rRNA gene amplicon data and the diversity of the samples obtained from the metagenomes presented in Thomas et al., (2014). This suggests that the microbial community composition in the subsurface of the Dead Sea is relatively independent of depth and salinity. Additionally this stands to support the reliability of our diversity data. It is most probable that similar microbial communities may arise from similar depositional environments, such as those shared by CT2 and M2. As described before, aragonitic facies were deposited from a stratified water column, in which fresh water inputs allow the massive development of specific organisms. Nutrient inputs by this freshwater, as well as biomass creation from blooming organisms may be reflected in the sediment. Carbon source may then be a second determinant factor for the microbial community composition. For example, Dunaliella bloomed in 1992 in the Dead Sea, after a dilution of the surface water down to 70% of its original salinity. Members of the Halobacteriaceae family were observed to bloom subsequently and were identified as the prime degraders of the organic matter produced by the algae (Oren, 1993). Therefore, the dilution of the upper layer by freshwater is responsible for shifts in microbial communities dynamics in the lake. This activity changes at the surface of the
38 Chapter II. Current microbial life in the Dead Sea sediment lake may well be transferred to the sediment, and induce differences in subsurface assemblages, between halite-gypsum sediment (from a holomictic phase) and aragonitic sediment (from a meromictic phase). TOC values cannot really infer a difference between aragonitic and true halitic facies. Few pure halite deposits were measured and gave values varying from 0.02 % for CT1 to 0.44 % for CT4. They are indeed lower than average, which is expected given higher sedimentation rate and little allochthonous input (low C/N values), which carries most of the organic matter to the Dead Sea (Oldenburg et al., 2000) in halite intervals. However, the quality of organic matter, which has been described as a key factor for microbial development in lacustrine sediments (Glombitza et al., 2013), could critically change with depth, and between sedimentary facies. The retrieval of KB1 and MSBL1 clades sequences could thus be linked to the specific setting of the lake at time of sedimentation.
2.2.4.4 Microbial community of halite-gypsum samples
Interestingly, almost no bacterial clones were obtained from halite and gypsum samples. In fact, in the deeper parts of the core, KB1 Candidate Division related sequences represent the only retrieved bacterial DNA. In the most surficial sample, bacterial sequences obtained from the core are those of moderate halophiles. Regardless of the possible biases induced by nested PCR, it is probable that Bacteria hardly subsist in such environments and are outcompeted by Archaea of the Halobacteriaceae family. The latter are Fig. 2.7 Cluster analysis using indeed favored by their highly energy unweighted pair-group average and efficient “high salt-in” osmotic Morisita index based on genera level of identified phylotypes. Cophenetic equilibration strategy (Oren, 2001). More correlation coefficient is 97.05%. than 65% of the identifiable OTUs in the
39 Chapter II. Current microbial life in the Dead Sea sediment core belong to this obligate halophilic family, which has already been described as the prime inhabitant of the present Dead Sea water column (Bodaker et al., 2010; Rhodes et al., 2012). Within this group, 91 OTUs have been attributed to the Halorhabdus genus, for which two species, H. utahensis and H. tiamatea, have been cultured in the past (Wainø et al., 2000 ; Antunes et al., 2008). Among the other representative sequences of the Halobacteriaceae family, some belong to yet uncultured Halobacteriaceae, most of which seem to be as well closely related to the Halorhabdus genus (Fig. 5). Members of the Halorhabdus genus grow both aerobically and anaerobically, and are able to use various substrates, making them relatively versatile colonizers for such extreme environments that are often poor in nutrients and labile organic matter (Werner et al., 2014). The Halobacteriaceae family is well suited for taking advantage of the variability of substrates like hydrocarbons, acetate or fatty acids, that may be available under the most extreme Dead Sea conditions (Nissenbaum, 1975; Oren, 1988). For example Halonotius pteroides has been evidenced to rely mostly on pyruvate, with ability to use glucose and glycerol as sole carbon source (Burns et al., 2010). Halomicrobium species H. mukohataei is able to decompose various sugars along with glycerol (Oren et al., 2002). Halobacterium strains previously isolated from the Dead Sea share common sugar fermentation abilities (Oren, 1983a). Haloplanus natans uses glucose, ribose and acetate as carbon and energy sources (Bardavid et al., 2007). And Haloterrigena salina uses various sugars and organic acids, notably acetate (Gutiérrez et al., 2008). We were not able to directly identify these species given the low similarity percentages between the Halobacteriaceae sequences we obtained and those in the databases. Nevertheless, the versatility over substrates used for fermentation is supported by metagenomic analysis of the Halobacteria-dominated gypsum sample CT3, where genes related to fermentation of sugar alcohols or amino and organic acids have indeed been identified (Thomas et al., 2014).
2.2.4.5 Microbial community of aragonitic samples
The only aragonitic sample in the core, for which clones could be obtained, is the shallowest one. To assess whether our method was efficient in such sediments, we inoculated aliquots of aragonitic samples from throughout the core with known DNA,
40 Chapter II. Current microbial life in the Dead Sea sediment which we were subsequently able to extract, amplify and clone using our protocol. We can infer that no enzyme inhibitor was preventing our protocol from working and that it is more likely that cell abundance was extremely low in the deeper aragonitic layers. The CT2 specific assemblage is largely dominated by Bacteria of the KB1 Candidate Division and by Archaea of the MSBL1 Candidate Division. The latter belongs to the Thermoplasmata class and has been defined in the hypersaline brines lying in the deep basins of the Mediterranean Sea (van der Wielen et al., 2005). The MSBL1 Candidate Division members are uniquely found in anoxic and hypersaline environments where sulfide is present and often where concentrations of divalent cations are very high (Antunes et al., 2011). In the deep Mediterranean anoxic basins, MSBL1 sequences were the dominant sequences in strata where methane was produced (van der Wielen et al., 2005; Borin et al., 2009). Based on their phylogenetic relationship (Fig. 5), their metabolism was tentatively related to methanogenesis. The KB1 Candidate Division has been identified in the hypersaline brines of the Kebrit Deep, in the Red Sea (Eder et al., 1999), a very similar environment to that in which the MSBL1 Archaea were found. Similarly, this brine is anaerobic, has a 26 % salinity, a high content of H2S and significant amounts of methane (Backer and Schoell, 1972). This group has since then been exclusively found in hypersaline anoxic environments, such as deep brine basins in the Mediterranean Sea (Borin et al., 2009; La Cono et al., 2011), or in hypersaline sediments like in the Lake Chaka sediments, on the Tibetan Plateau (Jiang et al., 2007). A recent study identified the association of MSBL1 and KB1 candidate divisions in the deep hypersaline methane-rich brine of Lake Medee (Yakimov et al., 2013) where it was able to establish that in such environments, the assemblage works as a consortium mostly relying on the degradation of osmoprotectant amino acids. While KB1 are thought to degrade glycine betaine (GB) and choline into trimethylamines and acetate, MSBL1 are performing methanogenesis from the methylated substrates resulting from this cleavage. Choline and GB are two common solutes used by halophilic organisms to deal with osmotic stress. This strategy is largely used by eukaryotes and Bacteria and is commonly named “salt-out” or “low salt-in”. It is more energy consuming than the “high salt-in” strategy used by Archaea of the Halobacteriaceae family, potentially explaining the large dominance of this family in the undiluted Dead Sea (Rhodes et al., 2012). While the “high salt-in” strategy dominates halite-gypsum samples (based on phylogenetic similarity), it appears that the aragonitic
41 Chapter II. Current microbial life in the Dead Sea sediment sample community could rely on the products of organisms using the “low salt-in” strategy, and more specifically on the osmoprotectant glycine-betaine. Metagenomics study of samples CT2 and CT3 show that genes for both types of osmotic adaptations are present in the microbial community of both samples. However, genes related to methanogenesis from methylated compounds are present only in the aragonitic sediment (Thomas et al., 2014) and subsequently support the existence of a cycle linked to the degradation of osmotic solutes. These osmotic solutes could originate from diluted waters from the epilimnion of the Dead Sea, potentially explaining the specific microbial assemblage of the aragonitic sediment, when compared with those of gypsum and halite. More work will help to be conclusive on such differences.
2.2.4.6 Microbial communities in continental sediments
While extensive geomicrobiological investigations have been carried out in oceanic sediments for more than two decades now, ranging from continental margins to abyssal sediments (Parkes et al., 1994; D’Hondt et al., 2009), very little is known on the microbial activity and diversity in lacustrine sediments. First geomicrobiological studies in the lacustrine realm show major variations of assemblages with depths and geographical locations, with a direct influence of the lake setting and local climatic changes. In Tibetan lake sediments, Dong et al. (2010) show that microbial abundance and diversity respond to different environmental gradients. In the sediments of a Patagonian maar lake, Vuillemin et al. (2013) emphasize that differences in lake productivity and organic matter preservation induced by climatic variations in the last 1500 years influence microbial communities throughout the sediment. In contrast, in the ocean subsurface, microbial communities organize along a general suite of metabolic processes (e.g. Froelich et al., 1979; Jørgensen, 1983), primarily determined by geological and geochemical characteristics of the sediment (Biddle et al., 2005; Inagaki et al., 2006). The great depth and slow response time of oceanic environments may thus prevent recording of climatic influence on microbial communities. The present work is a first attempt at characterizing both archaeal and bacterial communities using 16S rRNA gene sequences in the lacustrine subsurface of the Dead Sea. Finding these sequences is a first at those depths in such extreme environments allowing a scattered but representative view of their existence and sustainability under
42 Chapter II. Current microbial life in the Dead Sea sediment deep hypersaline conditions. It adds to the currently growing knowledge of continental microbial subsurface environments (Ariztegui et al., 2015). It also emphasizes the importance of implementing routine geomicrobiological investigations in the framework of future ICDP projects. Indeed, sediments of the Dead Sea and many other lacustrine environments are primarily used for their potential for high-resolution paleoclimatic reconstructions. This study suggests that even in extreme environments, a pre- evaluation of the potential proxy disturbance that can arise from microbial activity is compulsory. If well constrained, subsurface microbial communities may additionally be used for local limnological and paleoclimatic reconstructions. Along with sampling for DNA and cell counting, systematic analysis of chemical compounds involved in metabolic processes (such as methane, sulfate, nutrient or DOC) is recommended, although in the current study, the Dead Sea environment imposed unmanageable challenges with respect to some of the analyses. Nevertheless, we can truly attest to the specificity of subsurface microbes in the lacustrine realm, being highly influenced by terrestrial input induced by changes in climatic conditions at the time of deposition. As a result, the diversity and versatility of deep lacustrine communities challenges any consensual model.
2.2.5 Conclusion
This study has evidenced for the first time the existence of a deep subsurface life in the Dead Sea sediments. Although the quantity is low, and metabolic rates remain unknown, clear similarities were found between modern communities at the water sediment interface, and those extracted from 90 and 200 m below the lake floor in similar sedimentary facies. This call for a reconsideration of estimated turnover rates and activity in the deep hypersaline sediment. Different lithological types hold specific microbial populations representing diverse environments. Potentially, the quality of organic matter provided by the water column during periods of high freshwater input allows development of specific communities, which could not survive in current Dead Sea conditions. Members of KB1 and MSBL1 Candidate Divisions could take advantage of the osmotic solutes available in the aragonitic sediments. Halite and gypsum sediments are dominated by Halobacteriaceae, which are today the only inhabitants of the lake water column. Their potential for survival, while still not resolved, appears to be
43 Chapter II. Current microbial life in the Dead Sea sediment immense given their presence 200 m below the lake floor , and potentially active to at least 90 mblf, as suggested by the RNA-based CARD-FISH images. Although highly challenging, the chemical and sedimentological characteristics of the Dead Sea sediments have permitted the establishment of a unique relationship between microbes and their lacustrine environment. The similarity shared by gypsum-halite samples communities, and the importance of freshwater inputs for assemblages of the aragonitic sediment imply that the lake water balance at time of sedimentation is in part responsible for the current distribution of microbes in the sedimentary column. Conclusively, changes in the general climatic conditions over the Levantine area markedly influence the lake water and the microbial community in the sediment. High sedimentation rates and extreme physical and chemical processes taking place in the Dead Sea Basin help recording those changes which are still visible in today’s subsurface biosphere.
2.3 Fluid inclusions as time capsules of chemistry and life
Halite precipitates regularly from the lake water column as observed from the lake surface in December 2010 (Fig 2.9). Halite crystals nucleation and growth from hypersaline waters is well documented, in the Dead Sea (Gavrieli, 1997; Stiller et al., 1997) and in other systems (Lowenstein and Hardie, 1985; Schubel and Lowenstein, 1997). This crystallization being heterogeneous, formation of depression or cavities within the crystal lattice is common along the growth lines. They may trap water and even organisms at the moment of precipitation, and in this way form time capsules for the fluids from which halite precipitates. These fluid inclusions have been regularly used in paleoenvironmental reconstructions to measure changes in elemental and isotopic concentrations, as well as for physical parameters such as salinity and temperature. Indeed, the fluids trapped in the inclusions, if they remain in a closed system, record the physical parameters of the water of precipitation. Methods such as microthermometry use this closed system to study phase changes as a function of temperature. As an example, the temperature at which a two-phased fluid inclusion reaches homogenization has been shown to be the temperature of precipitation of the halite crystal (Lowenstein et al., 1998). This method has been tentatively used for the halite recovered from the ICDP cores. Protocoles and preliminary results are presented in
44 Chapter II. Current microbial life in the Dead Sea sediment appendix (Fig. A4-A5) and could be coupled to isotopic measurements of oxygen and deuterium to contrain paleotemperatures of interglacial periods in the Dead Sea Basin (Rigaudier et al., 2011). While fluid inclusions can trap fluids, they have also been evidenced to trap microbes present in such waters. In this manner, micro-organisms and their cellular material are preserved, and even arguably kept alive/viable in these tiny microenvironments (Norton and Grant, 1988; Vreeland et al., 2000; Lowenstein et al., 2011). Thus, they present a good potential for unravelling changes in microbial ecology during the history of hypersaline lakes. As todays microbiology techniques allow the study of supposed living sedimentary communities, micro-organisms trapped in the fluid inclusions would allow a priori comparison with organisms living in the water column at time of sedimentation. Regarding geomicrobiology hot topics, such as the origin of subsurface communities, it can also inform on the interactions between water column assemblages and sedimentary ones. Based on these observations, careful microscopic examination of fluid inclusions from halite intervals was done. Traces of organic matter, observed under visible and UV light (Fig. 2.9), as well as identification of Dunaliella algae in the inclusions (Fig. 2.10) confirmed the potential for preservation of cellular metabolites, and potentially of DNA within the Dead Sea fluid inclusions. Extraction of DNA and subsequent amplification and cloning of 16S rRNA gene was realized following a protocol tailored for such
Fig. 2.8: Few millimeters halite rafts precipitating at the surface of the Dead Sea in the late morning of December 2010. Photo credit: Aurèle Vuillemin.
45 Chapter II. Current microbial life in the Dead Sea sediment material and such little DNA quantity (Sankaranarayanan et al., 2011). The acid bleach protocole was used as follow: crystals of halite were first soaked in 10N HCl prepared at minimal halite solubility for 15 minutes. They were then transported to a neutralization solution of Na2CO3 10 % prepared at halite saturation and to 6 % NaOCl (at halite saturation), for 15 minutes for both. Rinsing in halite saturated brine was performed between each step, in order to avoid reagent carryover, and a final set of 4 washes of 15 minutes each, was done to secure the complete cleansing of the halite crystal walls. After dissolution to 1 M, samples are further desalted using Amicon Centrifugal filters (Ultra- 0.5 50 kDa, YM50, Millipore). The filters allow retaining only DNA fragments longer than 50 bp. These fragments were further purified using a minElute PCR Purification Kit (Qiagen). Environmental PCR and nested PCR followed similarly to what is presented in section 2.3.2, for both Bacteria and Archaea. Cloning and sequencing was performed and allowed the recovery of 13 different sequences from four different halite samples (Table 2.5). These fluid inclusions led to 8 archaeal sequences, all related to the halophilic Halobacteria. Bacterial sequences are related to Gammaproteobacteria of the Altermonadaceae family. It is potentially related to the halotolerant Marinobacter genus. At 32.9 m blf, bacterial sequences related to the Actinobacteria and the Firmicutes phyla were the only obtained sequences. The Firmicutes sequence is related to the Halanaerobiacae family, also described in the microbial mat aragonite lamina. The Actinobacteria sequence is related to the halotolerant Yaniella genera. In sample A-26, sequences related to Rhodovibrio, a photosynthetic halophilic Alphaproteobacteria and to the KB1 Candidate Division, the latter representing the only bacterial taxon of the sedimentary assemblages (aad facies at 2.74 mblf; section 2.3) are the only bacterial sequences of the 8 obtained sequences. The others relate to Halobacteriaceae putatively of the Halorhabdus genus, which is allegedly the most represented genera of the deep halite and gypsum sequences. Additionally, 3 sequences of the halophilic MSP41-clade were recovered. This clade was also obtained from the microbial mat. The deepest samples, from core A-42 (87.6 m blf) and A-82 (206.53 m blf), did not yield any sequence. The recovery of such sequences does not allow a full understanding of the factors contributing to microbial assemblage settling in the lake environment, nor does it permit to link water column communities with sedimentary ones. In situations of halite precipitation, the lake is expected to be in an oxic holomictic stage. Sequences attributed
46 Chapter II. Current microbial life in the Dead Sea sediment to anaerobic species such as Halanerobiaceae, or KB1 Candidate Division are not really expected cast some doubts regarding the complete sterilization of crystal surfaces. Indeed, although halite precipitation is not only restricted to the surface water (authigenic growth forming large crystals or spontaneous halite precipitation from the deep water participating actively in the formation of fine grained halite accounting for 50 % of the modern halite fraction have been evidenced (Gavrieli, 1997), these type of precipitation are very unlikely to form fluid inclusions and thus to preserve living cells in their lattice (Shearman, 1970; Lowenstein and Hardie, 1985). Moreover, these doubts are erased when comparing sequences obtained for example from sample A-1 between the fluid inclusions and the bulk sediment (CT1 in previous section), or from the absence of fluid inclusion sequences from sample CT4 at 206 m blf. Considering the recovery of sequence from its bulk sediment, and its reproducibility, the protocole seems to work. It was moreover successfully tested through contamination by allochthonous DNA, which could not be retrieved after the sterilization process. Explanation for the existence of such organisms in fluid inclusions could thus be that (A) sediment organisms can be transported to the water column and survive in aerobic conditions for a given amount of time; (B) conversely, sediment organisms originate from the water column and can be preserved in the water even when the latter has become oxic (Oren, 1983b; Bodaker et al., 2010); (C) halite growth continues at the bottom of the lake and even in the sediment where it can trap autochthonous communities. Continuous growth of halite in the subsurface has been acknowledged in salt pans, where specific “saw-tooth” along with fine grained halite facies are found (Gavrieli, 1997; Schubel and Lowenstein, 1997; Kiro, personal communication; Thomas, field observation). All hypotheses thus seem plausible. The second one is supported by the ability for residual community to remain in the water column for long period, even after the suppression of meromictic conditions which triggered their bloom (Bodaker et al., 2010). The advective transport of microbes from the sediment to the water column, suggested in the first hypothesis is not very likely as diffusion dominates and since advection has been ruled out by previous studies in the Dead Sea water sediment interface (Nishri, 1984). Information derived from the presence of Halobacteriaceae sequences, and more precisely from Halorhabdus genera
47 Chapter II. Current microbial life in the Dead Sea sediment
Fig. 2.9: Fluid inclusions from the Dead Sea core 5017-A. (A) Primary cubic fluid inclusions of the type used for microthermometry (see annex). (B) Visible black material, interpreted as a mix of detrital mud and organic matter trapped in halite crystals. Blurry dark areas are zones of high fluid inclusion density. Organic matter is not often visible under visible light (C) but can be clearly see through its auto-fluorescence under UV light (D). Scale bars are 50 µm except for (B).
Fig. 2.10: Putative Dunaliella cell enclosed in a fluid inclusion in a halite sample in core A2 (1.3 mblf) from the Dead Sea drill hole 5017-A.
48 Chapter II. Current microbial life in the Dead Sea sediment
Table 2.5 sequences recovered from fluid inclusions of the Dead Sea core samples are less decisive. Indeed, the presence of these highly adapted halophiles is recognized in the Dead Sea water column (Bodaker et al., 2010; Aharon Oren, 1983b) as well as in its sediment (Thomas et al., 2014). Halorhabdus members have been acknowledged to be highly versatile and display adaptations to oxic and anoxic environments, preventing any inference on their environmental niche. Sequences related to halotolerant species are not likely to develop in the halite-precipitating Dead Sea water column, salinity conditions presumably being to harsh for them (ad discussed in the previous section). They are more likely to have been transported to the lake before being trapped in the fluid inclusion. Finally, the recovery of a halophilic photosynthetic sequence of Rhodovibrio, underlines the potential of fluid inclusion extractions for trapping surficial water column communities to the sediment. Unfortunately, the poor coverage inherent to this method, and that of the sedimentary communities prevents conclusive statement regarding the origin of subsurface microbial communities. It only gives additional proofs of the diversity and complexity of the Dead Sea microbial ecosystems, in the sediment and the water column. Questions regarding the transport of microbes to the Dead Sea, and between its sedimentary and water pools are unsolved. The extent of activity of halotolerant and halophilic microbes must also be adressed before the current Dead Sea microbiome can be fully illuminated.
Acknowledgements
We thank Harald Gruber-Vodicka for his help with the phylogenetic comparison between the metagenomics data and the clone libraries. We also thank Aurèle Vuillemin, Nicolas Waldmann and Emanuela Reo for their contribution to the field and lab work, as well as Aymeric Le Cotonnec and Romain Vaucher for their help during sampling.
49 Chapter II. Current microbial life in the Dead Sea sediment
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Appendix material
Table A1. OTU definition and presence/absence in each sample
Table A2 OTU distributions and phylotypes definition
Table A3 Calibration values for microthermometry analysis
57 Chapter II. Current microbial life in the Dead Sea sediment
Table A4 Temperatures of homogenization for 4 samples of the Holocene part of the core
Fig. A1. Photograph of sediment sample incubated with tyramide showing non-specific
binding of the dye (green) on sediment
Fig. A2 16S rRNA gene based phylogenetic tree of the Bacteria representative sequences
for phylotypes defined in this study. Tree was built using the Weighted Neighbor
Joining method. Scale 10% of estimated difference in nucleotide sequences. Bars on
the right panel identify main bacterial classes and major clusters identifiable in the
Dead Sea sediments
Fig. A3 Comparison of microbial diversity in sample CT2 and CT3 obtained through 16S rRNA cloning (this study) and metagenomic data (A) Microbial diversity in the aragonitic sample. Metagenomes data from assembled 16S rRNA genes obtained from
AD metagenome presented in Thomas et al. (2014). (B) Comparison of archaeal diversity. Since metagenomic data did not provide convincing 16S rRNA gene fragments for sample GY (Thomas et al., 2014), comparison is made through total gene similarity with the M5NR database.
Fig. A4 Distribution of temperatures of homogenization of halite fluid inclusions
Fig. A5 Halite block used for calibration of microthermometry
58 Chapter III. Metabolic potential of microbial communities in the subsurface of the Dead Sea
“You can’t always get what you want”
Let it bleed, The Rolling Stones
59
60 3.1 Metagenomic studies in the Dead Sea subsurface
In order to gain insight into the microbial metabolisms that could occur in the Dead Sea sediment, two of the most interesting samples for which 16S rRNA gene sequences could be obtained (namely CT2 and CT3, see Chapter 2) were selected and sent for metagenomic analysis. The goal was first to obtain validation of the data obtained from our cloned amplicons. Although poorly known, results from the previous chapter suggested very low diversity with large dominance of the bacterial KB1 Candidate Division and the archaeal MSBL1 Candidate Division in CT2, and Halorhabdus-related genera in CT3. These samples are subsequently well suited for high-throughput sequencing method. They would allow a deep investigation of the metabolic potential of these relatively unknown communities. However, outputs from these methods revealed hard to interpret. Assembly was extremely complex and could not lead to reliable results. We thus decided to only present results linked to interpreted archaeal DNA, while continuing effort are pursued, using new assembly techniques such as that described by Albertsen et al. (2014), to properly reach bacterial communities, and potentially address the metabolic potential carried by the so far unsequenced Candidate Divisions MSBL1 and KB1 genomes. Issues pertaining to the environmental metagenomic approach in the Dead Sea subsurface will be presented in the next section.
3.2 Archaeal populations in two distinct sedimentary facies of the subsurface of the Dead Sea
A modified version of this section has been published in Marine Genomics as Thomas C., Ionescu D., Ariztegui D., and the DSDDP Scientific Team (2014) Archaeal populations in two distinct sedimentary facies of the subsurface of the Dead Sea, 17, 53-62. http://dx.doi.org/10.1016/j.margen.2014.09.001
Abstract
Two different sedimentary facies of the Dead Sea subsurface have been investigated for archaeal metabolisms using high-throughput DNA sequencing. We show that the communities are well adapted to the peculiar environment of the Dead Sea subsurface.
61 They are able to deal with osmotic pressure using high- and low-salt-in strategies, and can also cope with unusually high concentrations of heavy metals. Methanogenesis, for the first time identified in the Dead Sea is an important metabolism in the aragonite sediment. Fermentation of residual organic matter, probably performed by some members of the Halobacteria class is common to both aragonite and gypsum-rich sediments. The latter group represents more than 95% of the taxonomically identifiable archaeal metagenome of the gypsum sediment. Archaeal classes associated with sulfur reduction have also been revealed and are associated in the sediment with EPS degradation and Fe-S mineralization as revealed by SEM imaging. Overall, we show that distinct communities of Archaea are associated with the two different facies of the Dead Sea, and are adapted to the harsh chemistry of its subsurface, in different ways.
3.2.1 Introduction
The Dead Sea as we know it today is a remnant water body of the previous Neogene ingression of the Mediterranean Sea into the Jordan-Arava rift Valley (Zak, 1967). The so-called Sedom lagoon yielded numerous evaporite deposits in the area and finally disconnected from the sea. Waters originating from this lagoon have evolved within this framework, towards today’s well known Mg-Ca-Cl brines of the Dead Sea, due to interactions with the surrounding geology, and through various dissolution and evaporation processes (Zak, 1967; Stein et al., 2000; Katz and Starinsky, 2009). The Dead Sea is now lying at 427 m below sea level (2013) and its continuous retreat since 1978 (Abu Ghazleh et al., 2009) has led to an extreme salinity (TDS) of 348 g. L-1 (Oren and Gunde-Cimerman, 2012). In addition, its high chlorine and divalent cation concentrations (Cl- = 6.1 M, Mg2+=2 M and Ca2+=0.5 M ; Ionescu et al., 2012) make it harsher for microbes to develop (Oren, 2001), approaching MgCl concentrations of 2.3 M which are thought to be the upper limits for life (Hallsworth et al., 2007). While the Dead Sea chemistry is going towards abiotic conditions (Oren, 2010a), its history is different and life has been observed to thrive when sufficient water dissolution occurs. One such case is the outflow of submarine springs on the western shore of the lake, where a diverse microbial community has taken advantage of nutrient and salinity gradients to develop (Ionescu et al., 2012; Häusler et al., 2014). During events of rainy winters, water dilution in a mixed upper layer has also allowed blooms of the alga
62 Dunaliella and of its archaeal degraders of the Halobacteria class (Oren and Shilo, 1982a; Oren, 1983b; Oren, 2010a). During more arid periods, studies have shown that Dead Sea water column biota consists mainly of Archaea from the Euryarchaeota phylum, which are often seen as residual communities of periods of moderate salinity (Oren, 1993; Oren and Ventosa, 1999; Bodaker et al., 2010). These changes in lake physics and chemistry are thus influencing the biodiversity of the lake, as well as its geological record, through changes in authigenic precipitation of minerals. This is currently deeply investigated within the project of building a water connection between the Red Sea and the Dead Sea (Oren et al., 2004; Bardavid et al., 2007; Abu Qdais, 2008). In the framework of the paleoenvironmental investigation carried out within the ICDP- sponsored Dead Sea Deep Drilling Project (DSDDP), we were interested in recovering geomicrobiological information on the lake’s subsurface, particularly on the potential of microbes to interact with and influence geochemical proxies used for paleoenvironmental reconstructions. Emphasis was put on current subsurface communities, their adaptation to their surrounding and their potential metabolism in such hypersaline conditions. Investigations on Archaea communities in the water column and what influences their development have been regularly carried out (Bodaker et al., 2010; Oren, 1983b; Wilkansky, 1936). In contrast, there is currently no data on the contribution of these Archaea to the subsurface of the Dead Sea. However, in the marine subsurface (Biddle et al., 2006; Lipp et al., 2008) and continental saline (Waldron et al., 2007) sediments, they have been identified as major contributors to the biomass. Therefore, using metagenomic methods, we address here the metabolic potential and diversity of Archaea in the Dead Sea subsurface and their putative influence on two of the main sedimentary facies (Neugebauer et al., 2014): aragonite- detritus and gypsum (hereafter AD and GY, respectively). These sedimentary facies are linked to major changes in lake physics and chemistry, triggered by climatic changes or important meteorological events. We thus approach our data with an eye on the mechanisms influencing changes in communities, and its response to the water column influence.
63 3.2.2. Methods
The samples originate from ICDP core 5017-1A of the DSDDP expedition. This core was retrieved at coordinates N 31°30'28.98", E 35°28'15.60" (middle of the Dead Sea) at depth of 297 meters (one of the deepest point of the lake). Sample GY originates from a core catcher, at depth of 90.64 meters below lake floor. Sampling was done on the field in sterile conditions, in an onshore geobiology-specialized lab specially set up for the drilling expedition (December 2010) using autoclaved pre-cut syringes (see Vuillemin et al. (2010) for more details). Sample AD was taken from a core interval (2.74 m) during core opening party in June 2011 at GFZ Potsdam. Cores transported to the ICDP facilities at GFZ Potsdam were sawed in halves. One half was dedicated to picture and non- destructive measurements, while the other was saved for immediate sampling in sterile conditions using similar pre-cut syringes. To prevent any sampling of contaminated or oxidized parts, the syringe minicore was taken from the middle of the liner, and the surface part was removed. DNA was then further extracted using a phenol –chloroform extraction protocol modified from (Ionescu et al., 2009) Cells were extracted from 0.5 g of sediment after multiple cycles of PBS rinsing, 30 sec sonication and quick centrifugation. They were then incubated for 20 min at 95 °C in 0.5 mL lysis buffer of 0.1 M Tris, 50 mM EDTA, 100 mM NaCl and 1 % SDS (pH 8). 250 L of Phenol: chloroform: isoamyalcohol was added and the samples centrifuged atμ maximum speed after incubation for 10 min at room temperature. Supernatant was extracted again with the same process and the upper phase collected using Phase Lock Gel TM (PLG) tubes (5 Prime). It was then cleaned twice with 0.5 mL 24 :1 (v : v) chloroform : isoamylalcohol and DNA was precipitated overnight at -20 °C in 1 volume of isopropanol and 2 % (final volume) of 3 M sodium acetate (pH 5.5). Pellets were washed with 0.5 mL of 70 % ice cold ethanol after a 30 min centrifugation at maximum speed, dried and finally redissolved in 10 L molecular grade water. The QIAEX II Gel extraction Kit (Qiagen) was used to removeμ salt and purify DNA fragments according to manufacturer instructions. DNA extracts obtained from samples AD (aragonite alternating with detritus sample) and GY (gypsum sample) were sent to Mr DNA ™ lab, at Shallowater, Texas for whole genome amplification (through MDA) and metagenomic sequencing. DNA quantification was realized with the Qubit® dsDNA HS Assay Kit (Life Technologies) and amplification performed using the REPLI-g Midi Kit (Qiagen). Enzymatic
64 fragmentation was carried out for 150 ng of each sample using the Ion Xpress Plus gDNA Fragment Library Preparation Kit (Life Technologies). Fragments of 200-300 bp were obtained from the Ion Shear reaction after 8 min in a 37°C bath. After purification, Ion Xpress Barcode Adapters (Life Technologies) 13 and 14 were ligated to samples AD and GY respectively and underwent nick-repair following the manufacturer's instructions. Size selection at approximately 330 bp was done with the E-gel SizeSelect 2% Agarose Gel (Invitrogen). After new Qubit® determination, fragmented libraries were pooled to an equal DNA amount for each sample to create the final library. The latter was finally diluted to a concentration of approximately 78 pM, bound to Ion Sphere particles using the Ion OneTouchTM 200 Template Kit v2 DL resulting in 16.38% unenriched templated ISPs. After Ion OneTouch ES enrichment, template ISPs were sequenced on an Ion 318 chip using the Ion Torrent PGM (Life Technologies). Raw sequences were submitted to the MetaGenome Rapid Annotation using Subsystem Technology (MG-RAST) server for annotation and statistical analysis (Meyer et al., 2008). Annotations were made using the M5NR integrative database for proteins and M5RNA for 16S rRNA gene. Classification was made under MG-RAST default settings. Maximum e-values cut-off was 1e-5, minimum identity cut-off at 60% and minimum alignment length cut-off of 15. Functions were classified using the Subsystems approach supported by the SEED environment (Overbeek et al., 2005). Statistical values regarding sequence length were obtained from that platform, and by using FastQC. The metagenomes are publicly available on MG-RAST under reference IDs 4561562.3 and 4561566.3 for GY and AD respectively. Diversity indexes were calculated using PAST (Hammer et al., 2001) at the class and genus level, by excluding unclassified reads within the various phyla. Dominance D is
2 measured as D= ((ni/n) ) where ni is number of individuals of taxon i. Shannon-Weiner
Index H is H= ((nΣ i/n)ln(ni/n)) and evenness is taken as Buzas and Gibson’s evenness = H e /S, with S numberΣ of taxa. Scanning Electron Microscope investigation was done on samples after regular and critical point drying on a Jeol® JSM-7001 FA at the University of Geneva. Energy Dispersive X-ray spectroscopy was carried out on the same device. Samples were mounted on an aluminium stub with double-sided conductive carbon tape. An ultra-thin coating (15 nm) of gold was then deposited on the samples by low vacuum sputter coating.
65 3.2.3 Results
Over the two samples analyzed, 2147029 quality-controlled sequences were obtained. Among them, 1608197 sequences encoded for 1008583 peptides of which to 165818 at least one annotation was assigned. They have an mean length of 168 ± 63 bp and 171 ± 58 bp (for AD and GY respectively), with modal classes (of over 400000 sequences for AD and over 900000 for GY) of 230-239 bp for both samples. Overall, we obtained 128259 sequences attributed to the domain Archaea. The remaining sequences, (bacterial part) will be discussed elsewhere. Sample GY displayed higher number of sequences phylogenetically related to Archaea (77647) but smaller archaeal proportion than AD (20.9 % compared with 24.9 %, i.e. 50612 sequences for AD; Table 3.1; Table S1 in supp. material). Only few archaeal ribosomal rRNA gene sequences were obtained, therefore, classification using the M5RNA database produced few results. In sample AD the genera Methanobacteria, Methanomicrobia and Thermococci from the Euryarchaeota phylum were detected, and Nitrosopumilus from the Thaumarchaeota phylum. No archaeal SSU rRNA sequence could be retrieved from sample GY (Table 3.2). Therefore, hereafter, taxonomy for both samples was obtained from the closest relative in the database of annotated protein coding sequences as suggested by the MG-RAST server. Both samples presented almost similar number of archaeal classes. Diversity was much higher in sample AD compared to GY at the class level (Shannon index of 1.99 and 0.16 respectively. Sequences putatively affiliated to all of the archaeal phyla (Euryarchaeota, Crenarchaeota, Korarchaeota, Nanoarchaeota and Thaumarchaeota) were identified (Table 1). A small fraction of the sequences (0.01% for GY, and 3.1% for AD) could not be attributed to any specific phylum but were attributed to the domain Archaea. In both samples, members of the Euryarchaeota phylum were dominant, with 88.6 % and 99.1 % of the AD and GY sequences, respectively. In the latter sample 96 % of the Euryarcheota sequences were associated to the Halobacteria class, (resulting in a strong dominance index for GY (0.95). In the AD sample, no class was strongly dominating and the sequences were distributed between Halobacteria (25 %), Methanomicrobia (20 %), Methanobacteria (11 %), Methanococci (10 %), Thermococci (16 %), Archaeoglobi (8 %), Thermoplasmata (2 %) and Methanopyri (3 %) (Fig. 3.1), resulting in a larger evenness index (0.81 compared to 0.13 for GY; Table 2). Thermoprotei of the Crenarchaeota
66
Table 3.1: Sequence information and diversity indexes for both samples, calculated at the class level, for annotations against the M5RN database.
phylum covered 6.7 % and 7.1 % of the AD and GY sequences respectively. Unclassified Korarchaeota and Thaumarchaeota related sequences represented around 0.8 % and 0.7 % in the AD sample and 0.1 and 0.04 % in GY, respectively. No Nanoarchaeota-related sequences were detected in the GY sample while it formed the remaining 0.07 % of the assigned AD population. Classification extends only to the class level due to low coverage and is presented in the supplementary material S1. Function subsystems were annotated against the redundant M5NR database using the SEED classification. They correspond to three level of “functional roles” which classify the final function larger metabolic processes or structural complexes (Overbeek et al., 2005). Level-one subsystems have been put in quotes for clarity. This approach shows that all archaeal classes in sample AD with the exception of Nanoarchaeota appear to possess similar functions (Fig. 3.2). Variations are found in “iron acquisition”, “metabolism of aromatic compounds”, “motility and chemotaxis”, “nitrogen metabolism” (met.), “phosphorus met.”, “potassium met.” and “sulfur metabolism”, “stress response” and “virulence, disease and defense” subsystems (9 out of 27 obtained subsystems). For sample GY, functional information is unevenly distributed and is mainly held by Halobacteria. Highest sequence percentages were attributed to “amino acids and derivatives”, “carbohydrates”, “protein metabolism” and “clustering-based subsystems” (Fig. 3.2). In the “carbohydrates” subsystem, reads related to di- and oligosaccharides, sugars and alcohols and CO2 fixation are present. We identified a good amount of genes that are related to “stress response”, being either oxidative (309 reads for AD, 234 for GY), osmotic (149/17), nitrosative (151/0), in response to carbon starvation (0/19) or related to detoxification (6/0) and heat shock (239/0). “Membrane transport” genes, notably related to ABC transporters are significantly represented with 425 sequences for AD and 312 for GY. Genes related to synthesis and use of osmoprotectants such as
67
Table 3.2: M5RNA database (LSU and SSU rRNA fragments) annotations obtained for Archaea from sample AD.
choline-betaine uptake and biosynthesis were more numerous in AD than GY (113/17). It is also the case for trehalose uptake and utilization (11/76; AD/GY) and for biosynthesis of galacoglycans (278/73). Moreover, AD did display sequences related to glycerol uptake and utilization (133), which were not found in sample GY. Potassium homeostasis related sequences were obtained from AD (279) and GY (85) but surprisingly were not attributed to Halobacteria in the latter sample (Fig. 3.3A). In the “virulence, disease and defense” subsystem, most sequences were related to heavy metal resistance. Among them, AD had a greater amount of arsenic resistance and copper homeostasis related sequences (149/73 and 600/143 respectively), while cobalt-zinc- cadmium resistance and mercury resistance dominated in GY (20/111 and 0/14) (Fig. 3.3B). The main difference between the two samples is found on genes related to methanogenesis. Indeed, except for 2 sequences related to methanopterin biosynthesis (60 for AD), GY does not display any sequence related to methanogenesis strays (21 for AD), carbon monoxide induced hydrogenase (116 for AD), and one-carbon metabolism associated with methanogenesis (58 for AD) or methanogenesis from methylated compounds (8 for AD, related to Methanomicrobia; Fig. 3.4A). On the contrary, genes related to acetogenesis from pyruvate were present in both samples (500/132). Fermentation is also found in both samples (381 sequences for AD and 108 for GY), with specific lysine fermentation associated to the “amino acids and derivatives” subsystem (102/62) and anaerobic degradation of aromatic compounds (4/13; Fig. 3.4B). Electron- accepting and -donating reactions involved in fermentation are also highlighted through their enzyme related genes in GY and AD. This is the case for NADH dehydrogenase (0/4), glycerol 3-phosphate dehydrogenase (11/0), ADP-forming acetyl-CoA synthetase (229/1) and anaerobic respiratory reductases (60/0). Specific metabolisms are also of importance in our metagenomes. Namely, a good amount of reads related to “phosphorus metabolism” was obtained from sample AD and
68 GY (221 and 1082 respectively). They are related to alkylphosphonate utilization (1/12), high (85/1068; AD/GY) and low (30/2) affinity phosphate transporters and pyrophosphate-energized proton pump (101/0). “Nitrogen metabolism” related genes were also detected for both AD (422) and GY (362). Ammonia assimilation dominates metagenome AD (371) and is the only N metabolic pathway associated to GY (362). AD metagenome also comprises N fixation (42) and denitrification related genes (9) (Fig. 3.4C). Numerous genes from both metagenomes could be assigned to “sulfur metabolism”. For AD, 211 reads related to inorganic sulfur assimilation were recovered, and 559 for GY. Among the aragonitic sequences, 18 related to an enzyme involved in the dissimilatory pathway of sulfate reduction (sulfate adenylyltransferase,
AD
GY
Fig. 3.1: Archaeal classes obtained from the AD and GY samples based on the M5NR database
69 dissimilatory-type, EC 2.7.7.4), present either in Thermoplasmata, Thermoprotei or Thermococci members. Sample GY also had sequences related to organic sulfur assimilation (16 reads) while none were found for AD (Fig. 3.4D). All subsystem-based annotations associated to their archaeal classes can be found in Table S2 of the supplementary material. Scanning electron microscopy using secondary electron imaging coupled with energy dispersive X-ray analysis has outlined the occurrence of Fe-S mineralizations along the core. Euhedral octahedron of pyrite (Fig. 3.5A) and globular iron-sulfur minerals (Fig. 3.5B) have been observed in aragonitic samples. Native sulfur concretions are also present in numerous intervals of the core, sometimes with dense distribution (Fig. 3.5C). Additionally, occurrences of Fe-S minerals, found embedded in microbial biofilms (Fig. 3.6A) are revealed by back scattered electron imaging. Their morphologies resemble globular to aggregated micrometric mineralizations. The biofilm-like texture (EPS) is common in the AD sample, as shown by the infilling of a presumably allochthonous diatom (Fig. 3.6B).
Fig. 3.2: Heatmap of level 1 Subsystems of the SEED classification for the various classes of the two samples based on annotations against the M5NR database using MG-RAST. Abbreviations: CV cofactors, vitamins, prosthetic groups, pigments; PP phages, prophages, transposable elements, plasmids
70
3.2.4 Discussion
In order to appropriately interpret the metagenomic data resulting from the subsurface environment of the Dead Sea, we first discuss the issues pertaining to the method itself. We then highlight the main metabolic features by focusing on stress response and metabolic potential of the communities, with an emphasis on major issues faced by hypersaline communities in subsurface environments.
3.2.4.1 Metagenomics in poorly characterized environments
Archaea from the sample GY are largely dominated by Halobacteria. Their abundance increases by a 7-fold factor compared with the AD sample, and encompasses almost 97 % of the complete metagenome. Unfortunately, low coverage and copy number probably precludes recognition and blast of non-conserved 16 S rRNA gene sequences. Regarding the functions, subsystems are very poorly represented by other classes. While coverage is likely to be incomplete in such environments, there is little chance that Archaea other than Halobacteria are truly present and active in the gypsum layer. Members of this class are thus expected to take part in most of the metabolic activity there. Such analysis emphasizes the issue of DNA origin, which is often raised when working with sedimentary environments “Can one be certain that the DNA analyzed comes for active sedimentary communities and not dormant or dead ones?”. When it comes to the Dead Sea, it is even more relevant since high salinities tend to slow down DNA degradation by decreasing depurination rates (Lindahl, 1993). Transport of microbes from surrounding environment is probably an issue, and non-halotolerant organisms will certainly lyse when in contact with the Dead Sea water. The remaining organic material is thus very likely to end up sedimenting, and the genetic material it carries will probably be mixed with that of autochthonous organisms. Since we have little possibilities for deciphering autochthonous from allochthonous sequences, we have chosen on a first approach to focus on Archaea. Secondly, we must take additional care in interpreting functional information when it relates to phylogenetic classes not known to present adaptation for hypersaline environments. In this way, the presence of sequences linked to genes for osmolytes uptake and synthesis is a key point of our dataset, and allows better constrain
71 on the organisms susceptible of being active in the Dead Sea subsurface. Regarding activity, classic geomicrobiology methods tested under the Dead Sea chemistry have failed. Non-specific binding of fluorochromes used for FISH techniques have precluded its efficient use (Thomas, unpublished), and ATP-tester tested in the field and shown to work in sedimentary environment (Vuillemin et al., 2010) were inefficient at such salinity. Assumptions on the activity of microbes in the sediment can thus only be made through remnant of their activity such as EPS production and degradation observed under SEM (Fig. 3.6A-B), or occurrence of authigenic iron sulfide minerals, H2S gas escape and sulfur concretions (Fig. 3.5A-C). Another potential hint is the lack of genes
Fig. 3.3: Heatmap of level 3 subsystems for AD (left) and GY (right) related to A. osmotic adaptation and B. heavy metal tolerance annotated against the M5NR database. In brackets are always the first, and when existing, the second subsystem level name. Abbreviations: CBS clustering-based subsystems, CH carbohydrates, do di- and oligosaccharides, MT membrane transport, os osmotic stress, PM potassium metabolism, Res Resistance to antibiotics and toxic compounds, sa sugar alcohols, Se stress response, VD virulence, disease and defense.
72
related to dormancy or sporulation. But its absence can also well be related to poor database coverage of those functions, or to poorly-suited methods for DNA extraction from spore-forming microbes. Obviously, the Dead Sea subsurface environment is very poorly known and does present a vast amount of novel genomic material, which precludes precise binning, or at least lowersits quality. This issue in hypersaline environments has already been raised by López-López et al. (2013) whom could not recover the taxonomic assemblages previously found in anoxic mats of a solar saltern using clone libraries (López-López et al., 2010). Approaching these metagenomes with customized pipelines, for example by using differential coverage binning to unravel rare taxa (Albertsen et al., 2013) is a currently investigated possibility. However, constant effort for obtaining whole genomes of species thriving in hypersaline anoxic sediments or water column such as that of the Mediterranean deep hypersaline anoxic lakes must be sustained. The task is rendered difficult by the fact that some of the dominating classes in these environments have been proven to have acquired specific metabolism through lateral gene transfer (LGT). This is the case for Halobacterium, which may have acquired a subset of aerobic respiration genes of bacterial origin through several events of LGT (Kennedy et al., 2001). This type of event seems to be particularly important in the Dead Sea, where lateral gene transfer from a halophilic Archaea and thermo- philic Bacteria of the Thermotoga class has been found (Rhodes et al., 2010). This work emphasizes the fact that LGT events may occur between organisms from different extreme environments, giving a potential explanation to the numerous sequences in our metagenomes that were related to thermophiles. Abundant reads associated with the crenarchaeotal Thermoprotei, Korarchaeota, and from euryarchaeotal Thermococci, Thermoplasmata and Archaeoglobi classes suggest the presence of thermophilic to hyperthermophilic organisms in the Dead Sea subsurface environment. While water temperature is lukewarm, and may exceed 30°C at the surface, it rarely reaches 25°C at the bottom, and thus does not make a suitable set for the establishment of thermophilic species. The LGT event occurrence may explain some of it. In their work, Rhodes and coworkers (2010) also stress the potential for recovering rare, unexpected but functioning organisms in the Dead Sea environments. Other possibilities could be that the obtained proteins share similarities with that of species of thermophilic classes, without truly belonging to those thermophilic species. Moreover, since not all relatives
73
Fig. 3.4: Heatmap of level 3 subsystems for AD (left) and GY (right when present) related to A. methanogenesis, B. fermentation, C nitrogen metabolism and D. sulfur metabolism. Annotation against the M5NR database. For A and B, in brackets are the first, and when existing, the second subsystem level names. It is the first second and third for Glycerol- 3-phosphate dehydrogenase and ADP-forming acetyl CoA synthetase. For C and D, nitrogen metabolism and sulfur metabolism are level 1 subsystems, line headers are level 3 subsystems. In brackets is the specific function for organic sulfur assimilation. Level 2 subsystems are either identical to level 3 or absent in the M5NR classification. Abbreviations: 1C one-carbon metabolism; AA amino-acids and derivatives; CH carbohydrates; cc central carbohydrate metabolism; CV cofactors, vitamins, prosthetic groups, pigment; ear electron-accepting reactions; edr electron-donating reactions; fd formate dehydrogenase; Fe fermentation; MAC metabolism of aromatic compounds; pm2 pyrvuate metabolism 2; R respiration; rd respiratory dehydrogenase 1. ** function level.
74 of a same class are thermophilic, the obtained resolution and that pertaining to the database cannot allow for more precision for now. Another option is a gradual adaptation of thermophilic species (Jiang et al., 2007), which could have been transported to the sediment from hot springs existing notably in the region of En Qedem, although water rarely exceeds 50°C there. Under time span of dozens of thousands of years, gradual adaptation may be possible for some microbial taxa, with thermotolerance lowered to temperatures of around 35 °C, commonly observed in the core (unpublished data). This explanation does not hold for the younger aragonitic sediments though, in which those thermophilic groups are also encountered, and even more numerous. Transport from hot shallow springs may be possible but it could hardly account for such ratios, especially since the contribution of these springs into the Dead Sea microbial population has been acknowledged to be very limited (Ionescu et al., 2012) link. Additional contribution from deep hydrothermal springs related to the faulting of the Dead Sea Basin, that have been evidenced through seismic data (Niemi and Ben-Avraham, 1997) is not known to date.
3.2.4.2 Coping with the harsh Dead Sea conditions
3.2.4.2.1 Salinity
The two metagenomes present significant amount of sequences related to stress response. However, samples seem to differ in the responses that can be given to this stress. Indeed, the aragonitic sample (AD) has a higher number of sequences (relative and absolute as compared with GY) directly related to “osmotic stress”, probably inferring that in the environment representative of the aragonite sediment, specialization towards high salinity is not dominant, as compared with the gypsum sample, which is better adapted to salinity. The presence of higher number of sequences related to choline and betaine uptake and biosynthesis indicates that the low-salt-in strategy for osmotic adaptation (Oren, 2008) is prevalent in the AD sample. This is confirmed also by abundance of features related to trehalose uptake utilization, and biosynthesis of galactoglycans and related lipopolysaccharides. Additionally, Thermoprotei and Thermococci classes exhibit functions related to glycerol uptake and utilization which are not present in the gypsum sample (GY), suggesting that a large
75 array of solutes is at the disposition of Archaea in this facies for osmotic equilibration (Roesser and Müller, 2001a) in the aragonitic sample. Additionally, sequences related to the cell membrane bound ABC transporter were detected. These have been reported in the past to be activated under salt stress (Mukhopadhyay et al., 2006). While the low salt-in strategy is present in both samples, one can also find a high amount of sequences related to “potassium metabolism” for the two metagenomes, indicating the potential for K+-homeostasis both in gypsum and aragonitic facies. The high salt-in strategy used by members of the Halobacteria group (Oren, 2001) can thus be also employed in AD and GY. Potassium uptake and efflux systems have been acknowledged to be key systems in Halobacterium species NRC-1, used as a model for haloarchaea (Ng et al., 2000). The overview of subsystems functions reveals that although more susceptible to osmotic stress, the aragonite community may be able to use various osmolites to cope with salinity, making the low salt-in strategy a dominant one for its community (Oren, 1999). However, Halobacteria communities, known as halophiles “par excellence” (Oren, 2002) may use the more exergonic strategy (high salt-in) in this facies. In sample GY, dominated by halobacterial-related sequences, K+ homeostasis will probably be preferred although sugar and choline uptake may be possible.
3.2.4.2.2 Toxicity of metals
Several genes related to Halobacteria in sample GY, and more generally to Euryarchaeota in sample AD point towards the high adaptation of the archaeal community to heavy metals, which for most of them, are moderately (Cd, Zn) to highly (Co, Cu) enriched in the meromictic Dead Sea water column and its interstitial water (Nissenbaum, 1977). Arsenic, cobalt, zinc, cadmium and even mercury for sample AD are dealt with a subset of resistance protein, pump driving and transporting ATPase and ion reductase, which are part of the larger system for heavy metal resistance in prokaryotes, and particularly in Archaea (Nies, 2003). Copper homeostasis related sequences highlight the importance of dealing with this metal in the Dead Sea chemistry. Copper concentrations has been shown to be very high and to vary greatly with depth, location and season in the Dead Sea (Nissenbaum, 1977). The system for copper homeostasis mostly relies on copper translocating P-type ATPase which has been shown to be an efficient transporter of copper across the cytoplasmic membrane in E. Coli (Rensing and
76 Grass, 2003) and is present in several archaeal extremophilic copper-resistant Ferroplasma genomes (Baker-Austin et al., 2005). It is found for Euryarcheota for AD and in halobacterial-related sequences for GY. Overall, Halobacteria of sample GY seem to harbor a large amount of adaptation to the metals, generally enriched in the Dead Sea sediment. At smaller depths in the aragonitic community, adaptations are less numerous and focus on arsenic and copper.
Fig. 3.5: Indications of active S cycle in the sediment core. (A) Euhedral pyrite in an aragonite (needle) rich sediment (65 m blf). Scale bar is 100 nm; (B) Fe-S globule in between aragonite needles and diatom fragments (338 m blf). Scale bar is 1 µm; (C) Core picture of an interval of aragonite laminae alternating with detrital laminae (aad facies) with numerous sulfur concretions (white). The core is around 10 cm wide.
3.2.4.3 Coping with deep sedimentary environments
3.2.4.3.1 Fermentation
Fermentation is well represented in the two samples and implies the use of various sources, from carbohydrates to amino acids and aromatic compounds. Enzymes
77 involved in electron donating and accepting reactions in E. coli anaerobic respiration were also detected, such as NADH dehydrogenase for AD and GY, and anaerobic respiratory reductase for AD. Additionally, the specific archaeal enzyme involved in degradation of acetyl-CoA to acetate is abundant in sample AD, and only found once in sample GY. This ADP-forming acetyl-CoA synthetase is specific to Archaea employing fermentation (Schäfer et al., 1993). While fermentative activity cannot be attributed to any specific phylum in sample AD it appears that Halobacteria are the only ones harboring fermentation related sequences in sample GY. Generally, they are considered as the main aerobic heterotrophs of hypersaline environments (Oren, 2010b), but members of this class are also known to degrade organic matter in anoxic settings, similar to the Dead Sea sediment. This the case for Halorhabdus sp. (Wainø et al., 2000; Antunes et al., 2008) and Halobacterium sp. (Oren, 1983a). Here in sample GY, they are evidenced to be able to perform anaerobic degradation or aromatic compounds and lysine. The latter also potentially occurring in sample AD, but not attributable to a specific class or even phylum. Acetogenesis seems to be occurring in both samples too, as observed by detection of proteins related to acetogenesis from pyruvate. For both samples, fermentation of different products is possible. Halobacteria are major actors in both samples, although they would probably use different substrates: preferentially mixed acids in sample AD, and amino-acids, sugars and aromatic compounds in GY. In sample AD, they could be assisted in this degradation by Archaeoglobi, Thermococci and Thermoplasmata members mainly.
3.2.4.3.2 Methanogenesis
We bring data that strongly suggest that methanogenesis is dominant in the aragonitic sample and is probably a major process in comparison to the gypsum sample. This is highlighted by various methanogenesis strays detected for AD, carbon monoxide induced hydrogenase, which is a key enzyme involved in methane and acetogenic metabolism (López-López et al., 2013), and methanogenesis related 1-C metabolism hits. Additionally, pathways for methanopteryn biosynthesis, a cofactor in methanogenesis (van Beelen et al., 1984) were detected. Methanogenesis from methylated compounds is an important metabolism in the aragonitic sediment (Fig. 3.4A). Under extreme salinities, the substrate for methanogenesis is more likely to be a
78 methylated one, as more energy can be derived from its degradation and thus be directed towards osmotic adaptation (Oren, 2010b). The only sequences related to methanogenesis from methylated compound are attributed to putative members of the Thermoprotei and Methanomicrobia classes. Among the latter, Methanosarcina, which could be detected through their 16S rRNA gene signature, are notably able to perform methylotrophic methanogenesis (Garcia et al., 2000). This class also host the rare halophilic methanogenic genera such as Methanohalobium (Zhilina and Zavarzin, 1987) and Methanohalophilus (Paterek and Smith, 1988; Boone et al., 1993; Davidova et al., 1997). An in-depth study of the genomes of these organisms will give more precise information concerning identity of methanogens, substrate use, and adaptation to high salinity. Overall, the dominance of “carbohydrates” and “amino acids and derivatives” subsystems (Fig. 3.2) suggest an important role of heterotrophic metabolisms of sugars and amino acids in this sedimentary environment. Amino acids are required for growth and maintenance of the halophilic biomass, notably through the formation of compatible solutes for osmotic adaptation; they can also form sources of carbon for heterotrophic growth.
3.2.4.3.3 Sulfur reduction
Sequences related to the assimilation of inorganic sulfur are found in both metagenomes, however many more in sample GY than AD. The potential for organic sulfur assimilation (alkanesulfonate assimilation), was exclusively detected in sample GY. The latter can be used as an additional source for sulfur, as it has been highlighted in microbial mats of Pozas Azules (Breitbart et al., 2009). Alkanesulfonate assimilation is known to be expressed in E. coli in situations of S starvation (Eichhorn et al., 2000). This could be interpreted as a sign of good adaptation of gypsum communities to nutrient starving ecosystems. Interestingly, in sample GY, no specific dissimilatory-related enzyme related sequences is found while 18 have been in AD. Finding of sulfate adenylyltransferase of the dissimilatory type may thus support the occurrence of sulfate reduction in AD. In GY, sulfur metabolism may only relate to assimilatory sulfur reduction, attributed to Halobacteria members. Dissimilatory sulfate reduction has been only recently detected in the Dead Sea (Haeusler et al., unpublished), supporting several
79 geochemical analyses that have concluded its occurrence in the lake’s hypolimnion in the past. Accordingly, sulfate and sulfide S isotopes from the lower water mass before the 1979 turnover (Nissenbaum and Kaplan, 1976) or from the outcropping laminated and disseminated gypsum of the Lisan Formation (Torfstein et al., 2005), show signature of dissimilatory sulfate reduction in the stratified paleo-Dead Sea. Other indications observed in the DSDDP core, like common euhedral pyrite minerals (Fig. 3.5A), iron- sulfur globules (Fig. 3.5B), H2S degassing or native S° concretions (Fig. 3.5C) allow us to
Fig. 3.6: SEM pictures of active organic matter degradation and proofs of sulfate reduction in samples AD (A) Backscattered electron photograph of a biofilm in between euhedral halite and oxide minerals. Whitish spots are Fe-S mineralizations embedded within the microbial matrix. Their typical EDX spectrum is shown in the upper right corner. (B) Planktonic centric diatom exhibiting biofilm in its central depression. Arrows show putative microbes. Note potential gas vacuoles formed in the EPS (white arrows). Scale bars are 1 µm. suggest the presence of an active S cycle in the aragonitic sediments. In sample AD, SEM investigation under secondary and backscattered electron, coupled with energy. dispersive X-ray spectrometry shows the occurrence of Fe-S minerals, often embedded in a matrix which bares strong resemblance to microbial biofilms (Fig. 3.6A). The sulfide in Fe-S mineralization is likely to originate from the reduction of sulfate, available in the Dead Sea water (Nissenbaum, 1975a). Co-occurrence with microbial biofilms suggests active degradation of the available organic matter, potentially by fermenters and sulfate reducing microbes (Dupraz et al., 2004), which could lead to the accumulation of Fe-S minerals within its matrix. Production and particularly degradation of EPS, suggested by
80 microbes and gas vacuoles within the biofilm (Fig. 3.6B) outlines the in situ activity of the subsurface biosphere of the Dead Sea. Archaea of the two sedimentary facies display the potential for sulfate reduction, primarily based on inorganic sulfur. In AD, it can possibly be dissimilatory while too few information is available for GY. Generally imputed to Bacteria, it shows that alike in hot or acidic environments, Archaea may be able to use such mechanisms in hypersaline sediments.
3.2.4.3.4 Nitrogen cycle
Nitrogen cycling by microbial communities has been argued to be very limited in the Dead Sea (Stiller and Nissenbaum, 1999). Nevertheless, pathways towards the assimilation of ammonium, particularly abundant in the Dead Sea (Oren, 1983b), seem to be present in both metagenomes. Concomitantly, the recovery of Thaumarchaeota- related sequences, likely belonging to the Nitrosopumilus genus as highlighted by the SSU similarity (Table 2) supports the presence of nitrogen metabolism in this environment. However, this group is not likely to be adapted to high salinity and anoxic conditions. It only presents adaptations to highly oligotrophic environments (Pelve et al., 2011) and its representatives have been identified in suboxic settings (Labrenz et al., 2010). Additionally, sequences related to nitrogen fixation and denitrification were uniquely detected in sample AD, and attributed to Crenarchaeotea of the Thermoprotei class. This is probably imparted to the fact that nitrate availability is limited in the Dead Sea (Stiller and Nissenbaum, 1999). More work must be pursued in that direction in order to unravel metabolic potential of the subsurface communities of the Dead Sea regarding nitrogen.
3.2.5 Conclusion
Archaeal metagenomes recovered from two different sedimentary facies of the Dead Sea both show the impact of salinity on the microbial communities they host. Dominance of Halobacteria (notably in the deepest sample) and presence of numerous genes related to osmotic adaptation, either from the high- or low-salt-in strategy, highlight the harshness of this environment. However, the sedimentary environments of the Dead Sea are not
81 only extreme through their salinity since surviving organisms also have to deal with high concentration of heavy metals, nutrient starvation and shortage of high quality organic matter. Overall, the dominance of carbohydrates and amino acids subsystems suggest an important role of heterotrophic metabolisms of sugars in this sedimentary environment. While amino acids are required for growth and maintenance of the halophilic biomass, notably through the formation of compatible solutes for osmotic adaptation, they can also form sources of carbon for heterotrophic growth. Among them, methanogenesis from methylated compounds is an important metabolism in the aragonitic sediment. Methanogen-related DNA is for the first time shown to exist in the Dead Sea environment, validating pioneer works that predicted their occurrence (Marvin-DiPasquale et al., 1999). The gypsum sample is largely dominated by halobacterial DNA. In this layer, methanogenesis is inexistent, and most activity seems to rely on fermentation. Concordant proofs point towards an active sulfur cycle in the paleo-Dead Sea water column, and potentially in its sediment. Inorganic sulfur assimilation is possible on the basis of archaeal sequences observed in AD and GY metagenomes, with potential for dissimilatory reduction of sulfate in aragonite. Co- occurrence of Fe-S mineralization, native S° concretions and other traces of microbial activity confirm the role of microbes in the Dead Sea sulfur cycle. The complexity and poor knowledge of this type of environment is a clear barrier to the full understanding of the implication of these metagenomes. Complementing studies on these in this direction will help to better constrain the metabolism of such extreme assemblages. As an example, the complete assembly of the metagenomes of these samples, both from Archaea and Bacteria, will help unveil the specificity of such extreme environments, and the true metabolic potential of its inhabitants.
Acknowledgement
This research was funded by the Swiss National Science Foundation (projects 200021- 132529 and 200020-149221/1). The authors wish to thank A. Vuillemin for his support on the field and in the lab, and A. Martignier for her expertise on SEM analysis as well as two anonymous reviewers who greatly improved the quality of the manuscript.
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94 Zak I (1967) The geology of Mount Sedom. Thesis, Hebrew University of Jerusalem.
Zhilina TN and Zavarzin GA (1987) Methanohalobium evestigatus n. gen., n. sp.- An extremely halophilic methane-forming archebacterium(Methanohalobium evestigatus n. gen., n. sp.- Ekstremal’no-galofil'naia metanoobrazuiushchaia arkhebakteriia). In Akademiia Nauk SSSR, Doklady, pp. , 464–468.
Supplementary material available online at http://dx.doi.org/10.1016/j.margen.2014.09.001
Table S1: list of archaeal sequences with corresponding annotations to the genus level (M5NR database)
Table S2: list of subsystems sequences (M5NR database)
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96 Chapter IV. Influence of microbes on the sedimentary record of the Dead Sea Basin
“Everything is borrowed”
Everything is borrowed, The Streets
97 Chapter IV. Influence of microbes on the sedimentary record
98 Chapter IV. Influence of microbes on the sedimentary record
This section will be submitted under a modified form to the Depositional Record as Thomas C., Ebert Y., Kiro Y., Stein M., Ariztegui D., and the DSDDP Scientific Team, “Microbial sedimentary imprint on the Dead Sea sediment”
Abstract
A study of an ICDP core recovered from the middle of the modern Dead Sea has allowed the identification of microbial traces within this subsurface hypersaline environment. A comparison with an active microbial mat exhibiting similar evaporative processes allowed distinguishing characteristics of Fe-S mineralizations and exopolymeric substances (EPS) resulting from microbial activity. EPS were identified in the drilled sediment but unlike in other hypersaline environments, it appears that they have a limited effect on the precipitation of calcium carbonate in the sedimentary column. Sulfate reduction however plays a role in all type of evaporative facies, leading to the formation of diagenetic iron-sulphides in glacial and interglacial intervals. Their synthesis seems to occur under progressive sulfidation that generally stops at greigite because of incomplete sulfate reduction. The latter may be caused by a lack of suitable organic matter in this hypersaline, hence energy-demanding, environment. Pyrite may be found in periods of high productivity, when more labile organic matter is available. The carbon and sulfur cycles are thus truly influenced by microbial activity in the Dead Sea environment and this influence results in diagenetic transformations in the deep sediment.
4.1 Introduction
The Dead Sea is among the most saline lake in the world. It also has extremely high levels of divalent cations Ca2+ and Mg2+, making it even harder for life to cope with its chemistry (Bodaker et al., 2010). Few microbes have adapted to such environments. They are mainly Archaea members of the extreme halophilic class Halobacteria, as well as few halophilic Bacteria (Bodaker et al., 2010; Rhodes et al., 2012). In such environment, salinity gradients are salutary for life. Submarine freshwater springs host numerous and diverse microbial mats benefiting from the local dilution of the hypersaline brine (Ionescu et al., 2012). Rainy winters have also brought in the recent past, enough freshwater to the lake, to allow a 70% dilution of the shallow layers,
99 Chapter IV. Influence of microbes on the sedimentary record enabling blooms of the halophilic algae Dunaliella parva (Oren and Shilo, 1982; Oren et al., 1995). The highly labile organic matter produced by the autotrophic eukaryotes is immediately degraded by blooms of Halobacteria (Oren, 1983). Such gradients are also known to have occurred in the paleo Dead Sea and in its precursor Lake Lisan. The latter extended all the way to the Sea of Galilee (Lake Kinneret) as a meromictic lake exhibiting salinity-controlled stratification (Hazan et al., 2005). Evidences originating mainly from isotopic measurements from the deposits of this Lake Lisan, today exposed in the Dead Sea Basin, show that the sulfur and carbon cycles have been clearly impacted by life during the last glacial period (Torfstein et al., 2005; Kolodny et al., 2005). The recovery of a 457 m long core from the deepest part of the lake via the ICDP-funded Dead Sea Deep Drilling Project has shown a large potential for climate and paeloenvironmental reconstructions (Neugebauer et al., 2014; Lazar et al., 2014; Torfstein et al., 2015). It also provides a unique opportunity to explore the diagenetic processes occurring in the sediment of the Dead Sea, before they get exposed and fully desiccated. In order to constrain the impact of microbial activity on this pristine archive, a geomicrobiological investigation has been carried out. It revealed the importance of assessing the role of microbial activity in the early diagenesis of the Dead Sea sediment, highlighting differential distribution of microbes along the core, with diverse functional potential (Thomas et al., 2014; Ariztegui et al., 2015). We thus here intend to investigate the potential traces of past or current microbial activity in the Dead Sea depositional record, and qualify early diagenesis in its deep sediment. In order to do so, we have examined the sedimentology and mineralogy of an active microbial mat of the Dead Sea western shore, and compared the identified microbial traces with what could be observed in the cored material. This is put in the light of hypersaline sedimentary environments. Such environments often allow for the development of microbial mats, as their salinity prevents the development of grazing communities (Des Marais, 1995). Being often close to saturation with respect to halite, gypsum and calcium carbonates, it is often hard to decipher abiotic mineralization to organomineralization. The role of EPS as a matrix for mineral nucleation, particle binder or cation concentrator is a corner stone of the mineralization in hypersaline sediments (Dupraz et al., 2004; Dupraz and Visscher, 2005). We address its recognition and role in the Dead Sea realm, which is known to precipitate aragonite abiotically from the lake water column (Stein et al., 1997). We also take a deeper look into the Fe-S mineralizations recognized previously as diagenetic
100 Chapter IV. Influence of microbes on the sedimentary record phases from paleomagnetic studies (Ron et al., 2006). Studies of the subsurface waters of the Dead Sea Basin have highlighted major redox transformations affecting carbon, iron and sulfur among other (Avrahamov et al., 2010; Kiro et al., 2013; Avrahamov et al., 2014). It seems critical to assess the extent of such changes, and in particular observe if it reaches the deepest region of the Dead Sea Basin, or if it constrained to the aquifer/brine interface at its shoulders.
4.2 Geological setting
The Dead Sea is located on the lowest spot on Earth, at a current elevation of 427 m below sea level (2013). Intense use of the water of its catchment area, notably for irrigation and potash mining has led to a decrease in water level in the last 40 years at an average of 0.7 m per year (Abu Ghazleh et al., 2009; Tahal Group and Israel Geological Survey, 2011). Today’s water salinity is among the highest on Earth for a lake of this size, reaching 348 g.L-1 of total dissolved salts (Oren and Gunde-Cimerman, 2012). The Jordan River constitutes the main source of freshwater, nutrient and carbonate ions to the lake. In rainy periods, flash floods fed by canyons (wadis) cutting through the Judean Mountains and the Jordanian Plateaus are an additional source of freshwater. Before 1979, sufficiently rainy conditions allowed a diluted epilimnion to establish though the mixing of freshwater with the Ca-Cl brine of the Dead Sea. This meromictic situation allowed the setting of anoxic conditions in the hypolimnion. Anaerobic degradation of organic matter by sulfate reducing organisms has in such situations been invoked to explain the high H2S concentration and Fe-S occurrence in the hypolimnion (Nishri and Stiller, 1984; Nissenbaum, 1975). The latter disappeared quickly from the water column after complete overturn in winter 1978-1979 (Nishri and Stiller, 1984). Exceptionally rainy winters have led in 1980 and 1991 to the renewed formation of temporal meromictic conditions. This diluted epilimnion constituted a less extreme environment in which the halophilic Green Algae Dunaliella can develop and bloom, imparting a red color to the lake (Oren and Shilo, 1982; Oren et al., 1995). An array of evidences suggest the presence of relatively similar conditions supporting the influence of life on the geochemical record of the Lake Lisan sediment. For example, a general increase of the ∂13C of the aragonite laminae deposited during the last glacial period, as observed from the Lisan Formation in the Perazim Valley has been interpreted as the result of increasing autotrophic activity in its paleo-epilimnion (Kolodny et al., 2005). Sulfur
101 Chapter IV. Influence of microbes on the sedimentary record isotopes from gypsum intervals also support the occurrence of active sulfate reduction in the water column of the lake (Torfstein et al., 2005; Torfstein et al., 2008), presumably at the interface between its oxic diluted epilimnion, and its anoxic hypersaline hypolimnion. Sulfur concretions from the Masada section bear isotopic signatures interpreted as the result of incomplete microbial sulfate reduction (Bishop et al., 2013). The two limnological regimes of the lake are also reflected in the nature of the evaporitic minerals precipitating in both situations. During meromictic conditions, aragonite precipitation dominates (Stein et al., 1997). This is emphasized by alternating aragonitic and detritus facies, which is representative of glacial periods in outcrops and in the core material (Stein et al., 1997; Neugebauer et al., 2014). During periods of increasing aridity, the lake becomes holomictic and gypsum precipitation occurs, followed by halite deposition. This type of deposit, intercalated by detrital sediment is largely dominating in interglacial periods. The core material clearly shows such distribution, relevant to the study of microbial distribution in the lake. Since carbon and sulfur cycles greatly vary from one situation to another, we target our sedimentological study on these different types of facies in order to tackle the microbial impact on the Dead Sea sediment.
4.3 Material
The microbial mat was collected on the western shore of the Dead Sea, near the En Qedem spring site (Fig. 4.1B). It consists of a gypsum and halite-rich biofilm (Fig. 4.1E) colored by red pigments originating mainly from Dunaliella cells, which have developed in the diluted water of the ephemeral pond Fig. 4.1D. Microbial communities were analyzed (Thomas et al., Geobiology in review) and are composed of allegedly heterotrophic Archaea of the Halobacteria class mainly, and almost no bacterial sequence could be sequenced. Aragonite stellate clusters are present in the mat too as a result of aragonite needles precipitation from the pond water (Fig. 4.2A). Below this EPS-rich red layer (Fig. 4.2B) pigmented by Dunaliella algae carotenoids (Fig. 4.2C), alternating laminae of pure aragonite, mixed aragonite and detrital material, and detrital sediment only are found (Fig. 4.2D). They form subcentimetric alternations of typical uncompacted AAD. The aragonite lamina immediately underlying the mat crust host similar archaeal community based on the extracted 16S rRNA gene sequences (see Chapter II).
102 Chapter IV. Influence of microbes on the sedimentary record
Fig. 4.1: Overview of sampling sites and material. (A) Map of the Levantine region with Dead Sea location. (B) Location of drilling site 5017-1A (green square) and of the Qedem area, where the microbial mat was sampled (red square). (C) View of the ICDP-drilling platform at site 5017-1A. (D) Hypersaline pond north of Ein Qedem, where the microbial mat sample was taken. The pool is ca. 15m long. (E) Photograph of the mat (red, orange and green in color) with underlying laminated sediments (grey to white).
However, the bacterial community seems more diverse, with halophilic fermenters such as KBI Candidate Division and Halanaerobiaceae, and a sequence associated to Desulfohalobium, a halophilic sulfate reducer. A cubical section of sediment was cut with sterile material, immediately wrapped in Parafilm® M and stored at -4°C. Subsampling was done using sterile scalpel and picks for various microscopy analysis in the geomicrobiology lab at the Department of Earth Sciences, University of Geneva. Salinity of the pond has reached 36.9%, with concentrations of major elements as follow: Na+ = 2.12 M, Ca2+ = 0.58 M, Mg2+= 1.59 M
103 Chapter IV. Influence of microbes on the sedimentary record
- 2- and Cl =6.11 M and SO4 = 18.9 mM, slightly above those of the current Dead Sea water (Ionescu et al., 2012). The core material was taken from ICDP core 5017-1A of the DSDDP expedition. This core has been retrieved in November-December 2010 from the middle of the Dead Sea (N 31°30'28.98", E 35°28'15.60") at depth of 297 meters. All material was sampled onshore from core-catchers at a specially-tailored geomicrobiology lab, at Ein Gedi, Israel, preserving sterile conditions, except for the 2.74 m sample, which was taken during the core-opening party at GFZ Potsdam, Germany in June 2011, from core 5017- 1-2H-2. In all cases the outermost part of the samples were discharged in order to avoid oxidation effects and contamination with drilling or any other fluids.
Fig. 4.2: (A) Aragonite needles and Dunaliella (orange circles) in the water of the ephemeral pond in Ein Qedem ; (B) Photograph of EPS in the gypsum-halite mat crust ; (C) Close-up view of Dunaliella algae. Note that the nucleus is visible; (D) section of the mat and underlying sediment exhibiting the AAD behavior typical of the Lisan Formation.
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4.4 Methods
Samples were examined using an optical microscope on smear slides and polished thin sections (for halite rich samples). Scanning electron microscopy was performed after regular and critical point drying on a Jeol® JSM-7001 FA at the University of Geneva. Samples were then mounted on an aluminium stub with double-sided conductive carbon tape and gold-coated (15 nm) by low vacuum sputter coating. Cryo-SEM was performed at C-SEM, in Neuchâtel, Switzerland. It consists of a Philips XL30 ESEM –FEG equipped with a Gatan Alto cryo-transfer system. The microscope was operated at 20 kV under a pressure inferior to 10-4 mbar. The sample was freeze-dried in liquid nitrogen, transferred to the cryo-chamber stabilized at -140 °C, where it was sectioned using a blade, sublimated and sputtered with platinum. Energy dispersive X-ray spectrometry and back-scattered electron microscopy were used for chemical analysis for both SEM and cryo-SEM methods, allowing semi-quantitative determination of mineral chemistry. Mapping of different elements of the mat section was done using µXRF at the University of Geneva under an EAGLE µProbe machine with a rhodium tube at a voltage of 40 kV and a current of 420 µA with a 50 µm spot. Mineral distribution was then interpreted based on each image and relative color intensity using the software of the Adobe® Collection Suite and the Vision32® software version 4.953. Elemental composition of core 5017 1A was measured using XRF at GFZ Potsdam, Germany on an ITRAX corescanner of COX Analytical Systems. A chromium tube was used at 30 kV voltage and 30 mA intensity. These data only serve for qualitative interpretations as they originate from raw counts (in counts per seconds) with sufficient counting statistics from un- calibrated quick core sections scans as detailed by Neugebauer et al. (2014).
4.5 Results
4.5.1 X-ray fluorescence scanning
The microbial mat and its underlying sediment is composed of similar facies as those defined in the cores (Neugebauer et al., 2014). Halite mixed with gypsum resembles that of interglacial periods, except it is completely embedded in EPS, while aragonitic laminae alternating with detritus show identical distribution as those of the AAD facies. Their chemical signatures are highlighted by key elements (Fig. 4.3). Chlorine is present
105 Chapter IV. Influence of microbes on the sedimentary record in high and homogenous concentration in the sediment and halite (NaCl) precipitated on the surface of the section (topography image) as the sample dried while being mapped at high resolution. Sulfur is concentrated in two laminae, and shows good correspondence with strongest peaks of calcium, indicating gypsum minerals occurrence. Calcium (green) is present relatively homogeneously in the sediment just like chlorine, as it is highly concentrated in the Dead Sea water. Such high concentrations prevent from visualizing aragonite laminae. Higher fluorescence corresponds to gypsum minerals. Aragonite is best mapped with strontium, showing the presence of aragonitic precipitation in greyish to whitish laminae of the mat (light blue in µXRF). It is intercalated by detrital input mapped under iron bands (light yellow). Diffuse detritus are also present below this band, imparting to the aragonite laminae its greyish color. The “Fe x S” mapping allows highlighting the largest Fe-S mineralizations. Although some background noise is observed due to the superposition of a gypsum and detritus signature, higher fluorescence spots Fe-S minerals, under the form of pyrite, greigite or other transition phases and is represented in blue in the sketch (Fig. 4.3). Strongest Fe-S intensity is observed at the margins of the detrital lamina, within the aragonite laminae, and relatively few within the iron-rich detritus lamina. In the core, detailed XRF profiles highlight the lithologies and associated facies and are presented in Neugebauer et al. (2014). Here we add data of “Fe x S” over the S profile of core 1A, in order to emphasize Fe-S mineralization occurrences (Fig. 4.4). Signatures of sulfur concretions are also overlapping as resolution given by the figure cannot allow centimeter scale distinction. Their occurrences are highlighted in the core section. While S peaks in gypsum related levels and S-concretion intervals, Fe-S mineralization are correlated with smaller scale fluorescence of sulfur. Higher Fe-S peaks are found between 0 and 80 m, especially within the 45 to 75 m interval where Fe-S profile does not always correlate with S profiling, hence indicating other signatures than gypsum and native sulfur concretions. Within the 90 m gypsum interval, Fe-S is also peaking although intensity is lower. In general, intensity is lower for Fe-S profiles below 100 m, regardless of S fluorescence, but does imply general occurrence of Fe-S mineralizations all along the lower part of the core, regardless of the lithology. Examples of the morphologies and sizes of iron sulfide minerals are presented on the right side of Fig. 4.4, together with occurrences observed under different screening techniques, every 3 meters.
106 Chapter IV. Influence of microbes on the sedimentary record
Fig. 4.3: µ-XRF mapping of a microbial mat and the laminated sections of modern sediments along the Dead Sea shore. The dominating mineral is authigenic aragonite (Sr- mapping) with some gypsum (S and Ca). The darker layer is detritic material (calcite and clays). Disseminated Fe-sulphide is outlined by the «Fe x S» mapping, and is principally found in aragonitic layers. Star shaped crystals are the result of secondary precipitation caused by desiccation of the samples during measurement. See methods for details on the interpreted sketch. Scale bar is the same for each scanned image and is 2.5 mm.
4.5.2 Scanning electron microscopy
The mat section is concentrated with Mg, Cl and Ca, as observed in µXRF mapping and reprecipitation of NaCl during measurements. Drying artifacts must thus be carefully considered when observing minerals and putative organic structures in the Dead Sea. Indeed, precipitation may occur upon drying and organic structures will be overloaded with Ca, Mg and Cl ions when measured with EDX methods. Fe-S mineralizations are found under different chemistry and morphologies, and at various depths. Euhedral minerals forming octahedron have been found at depth of 65.38 m (Fig. 4.5A) in “laminated detritus” facies (ld, Neugebauer et al., 2014). Micrometer-scale spheroids were also found in such facies and also at 90.64 m in gypsum intervals (Fig. 4.5B), and rough-surface spherulites of ca 5 µm of diameter were retrieved at 337.62 m in “alternating aragonite and detritus” facies (AAD; Fig. 4.5C). Euhedral morphologies of 5 to 20 µm-scale pyrite were observed at the boundary between detrital and aragonitic
107 Chapter IV. Influence of microbes on the sedimentary record
Fig. 4.4: interpreted lithological profile of core 1A taken from Neugebauer et al., 2014, opaques, Fe-S mineralizations and EPS occurrences, and corresponding S and Fe S profile as given by XRF scanning. On the right hand, SEM photographs of (A) micrometric euhedral Fe-S mineralization from core 1-A-32 at 65.38 m; (B) Fe-S spheroid at 90,64 m, core 1-A-43; (C) rough-surface spherulite from core 1-A-131 at 337.62 m. Depth in meters below lake floor.
levels of the microbial mat (Fig. 4.5A). In the core they never reach such sizes and are generally individual euhedra with geometrical overgrowth of 5 µm maximum (Fig. 4.5B). True framboidal pyrites were observed in the mat (Fig. 4.5C) and seem to consist of small euhedral octahedrons. In the core, they are hardly recognizable and are generally formed of a limited amount of Fe-S spheroids (Fig. 4.5D), rarely exceeding 1 µm. They sometimes occur under the form of aggregated euhedral Fe-S precipitates (Fig. 4.5E-F). Size is also limited to a few micrometers of diameter. This form may be a precursor phase of micrometer-scale spherules or larger rough-surface spherulites (Fig. 4.5G and Fig. 4.5H, respectively). In the core, these precipitates rarely reach the pyrite stoichiometric composition (Fig. 4.6). The largest minerals (> 5 µm) approach true pyritic compositions (FeS2), being either euhedral minerals in the mat, or rough-surface spherulites at 350 m in the core. Additionally, in some dark high magnetic susceptibility laminae associated with AAD (Fig. 4.7A), Fe-S rims composed of greigite under were
108 Chapter IV. Influence of microbes on the sedimentary record found around Ti-magnetite detrital. In such settings, Ti-magnetite often present marks of dissolution (Ebert, personal communication). Although not fully characterized, Fe-S mineral phases occur around gypsum in detrital-rich halite intervals. They were never found when no detrital component was associated with the halite (Kiro, personal communication). Most other minerals, generally under the form of spherules or octahedrons stand close to the greigite chemistry line (Fe3S4). Few Fe-S precipitates lie below the mackinawite line (Fe8S9), and they correspond to sub-micrometric minerals. The EDX spot for measurements being > 1µM in depth and width, data from the smaller minerals should be taken with caution and have been highlighted in the Fig. 4.6. Exopolymeric substances have been clearly observed in the mat, under the naked eye, a regular microscope (Fig. 4.2) and cryo-SEM. Their cerebroid structure under cryo-SEM allowed a good determination in the mat (Fig. 4.8A). It extends in the whole mat sample, and displays a transparent film-like texture in close-up view, in which aragonite stellate clusters are often found embedded (Fig. 4.8B). The film may also be very smooth and present an apparent transparency under regular SEM, covering detrital and autochthonous precipitates (Fig. 4.8C). These characteristics allowed determining the presence of EPS in the core sediment. Indeed, chemistry cannot allow for clear distinction as the classical EPS carbon and oxygen composition is masked by Ca, Mg and Cl, highly concentrated in the Dead Sea environment. Using morphological identification by cryo-SEM allowed recognizing homologous sheet-like texture representing EPS in the core. The EPS are characterized by a film texture, sometimes folding on itself, or being torn apart during the preparation (Fig. 4.8D), possibly presenting voids and more rugged surfaces. Aragonitic stellate clusters or individual needles are often found embedded in these EPS. Cryo-SEM observations of the mat sample allowed to distinguish sharp freshly precipitated aragonite stellate clusters embedded in the cerebroid texture typical of the EPS (Fig. 4.9A). The thin needles constituting the stellate cluster are individually covered by the biofilm as observed in close-up view (Fig. 4.9B). These rough surfaces on the torn EPS are formed by nanoglobules (Fig. 4.9C). They are also observable in-between thin aragonitic needles, apparently entangled in this biological structure (Fig. 4.9D). Needles remain here very thin and elongated, without sharp tips and smooth surface. In the core, sharp and thin aragonite stellate clusters, as well as broken up thick ones may emerge or pierce through the EPS (Fig. 4.9E). They also are found aggregated
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110 Chapter IV. Influence of microbes on the sedimentary record
Fig. 4.5: Fe-S mineralization morphologies in the Dead Sea realm. (A) Euhedral pyrite from a white aragonitic lamina of the mat. Aragonite needles are visible in the background. (B) Similar euhedral morphology of Fe-S mineral in core 1-A-32, at 65.38 mblf. (C) Cryo-SEM picture of a framboidal pyrite embedded in EPS at the mat-sediment interface of the mat sample. (D) Agglomerate of four framboidal mineralizations in the core (65.38 mblf). (E) back-scattered electron imaging of a Fe-S rough surface spherulite being formed in core 1-A-30, at 59.29 mblf. (F) close-up view of (E) in secondary electron imaging, emphasizing the rough droplet like topography of the mineralization. (G) micrometer-scale Fe-S spheroid at 90.64 mblf, core 1-A-43. (H) Rough surface spherulite of micrometric scale at the tip of aragonitic stellate cluster in core 1-A-131, at 337.62 mblf. Pictures were all taken under secondary electron imaging in a regular SEM, except when specifically stated.
on its surface (Fig. 4.9F), or in between them and gypsum minerals, where they host numerous Fe-S minerals, as evidenced by back-scattered imaging (Fig. 4.9G). At larger depth (i.e. 350 m), EPS-like structures could not be identified although smooth film-like textures were found associated with aragonitic stellate clusters (Fig. 4.9H). Here, they correspond to halite minerals, demonstrating the complexity of untangling biotic features from abiotic ones in the Dead Sea sediment.
4.6 Discussion
4.6.1 Occurrence of traces of microbial activity in the Dead Sea sediment
EPS
The mat shows high production of EPS (Fig. 4.2B and 2D), also recognized unequivocally in the core. The recognition of EPS morphologies in the mat, through cryo-SEM and comparison with regular SEM structures and composition has allowed to distinguish them. Their recognition is generally complex in the wet Dead Sea sediment. Numerous poorly investigated precipitates, occurring in the Dead Sea naturally or upon desiccation before SEM observation, may take smooth sheet-like structure (Fig. 4.9H) similar to documented textures for EPS (e.g. Spadafora et al., 2010). Chemical analysis of hypothesized EPS is not equivocal as X-ray penetration from EDS generally exceeds the EPS thickness. Loading of these structures with calcium, magnesium and chloride has also tendency to mask its carbon and oxygen content. Morphological characterization
111 Chapter IV. Influence of microbes on the sedimentary record
Fig. 4.6: EDX composition of Fe-S mineralization in the Dead Sea subsurface. Pyrite line as y=1/2(x), greigite line as y=3/4(x) and mackinawite line as y=9/8(x).
was often the most reliable solution. Comparison with chemical composition and morphologies under the cryo-SEM allowed its conclusive identification. In the investigated sediment of the core, EPS detection always appeared in aragonite (AAD) or detritus-rich sediments. It was only very rarely that EPS could be identified in association with gypsum or halite minerals only. Several reasons can emerge from this observation. First, EPS recognition is harder in halite rich sediments, where smooth structures sometimes resembling that of EPS are attributed to amorphous precursors of various salts, potentially formed during sample preparation. Another reason is the enhanced preservation or production, occurring in the different sedimentary facies of the Dead Sea. Thomas and colleagues (2014) evidenced variations in metabolic potential in aragonitic and gypsum sediments. Organisms detected in these sediments may make use of EPS as C source, or protective matrix, given the adaptations to hypersalinity they harbor, suggesting preferential degradation or preservation in a given facies.
112 Chapter IV. Influence of microbes on the sedimentary record
Protection from the high Ca2+ and Mg2+ is probably the main reason for EPS production in the Dead Sea. The bonding capacity of EPS towards divalent cations (Kawaguchi and Decho, 2002; Dupraz and Visscher, 2005) could lower in situ concentrations of these cations, in which resides the main toxicity of the Dead Sea chemistry (Nissenbaum, 1975b; Bodaker et al., 2010). EDS measurements on EPS (unpublished) show a large concentration of calcium and magnesium in the EPS matrix. In the core, EPS are also observed. However, their occurrence is constrained mainly to the top of the core. Below that, they could hardly been recognized, suggesting the active degradation or difference/absence of production. The finding of EPS in the core shows that microbial activity has occurred at one point in the sediment. Their further disappearance may also support anaerobic heterotrophy.
Fe-S minerals
In the mat, several types of iron-sulfur minerals were observed. Framboidal pyrites were located close to the microbial mat only while large euhedral pyrites were observed in the underlying aragonitic laminae (Fig. 4.3). In the core however, although euhedral morphologies were observed, mineralizations were generally small (<2 um diameter). Their chemistry is also significantly different, with Fe/S ratio limited to that of mackinawite or greigite (Fig. 4.6), and very few pyrite. The microbial mat revealed to be of great help to allow identification of mineral phases in the Dead Sea environment. As clearly exhibiting biological processes, careful examination of the mat layers allowed the identification of morphologies in an environment characterized by microbial activity in the Dead Sea chemistry. Linking morphologies with processes of formation is still a matter of debate. Composition and morphologies result of numerous parameters such as the variety of reactive iron available, the saturation rate, the concentration of reduced sulfur and stability of redox conditions (Berner, 1984; Wang and Morse, 1996; Wilkin and Barnes, 1997; Wilkin and Arthur, 2001). Combination of observations from field and laboratory models attempting to mimic sedimentary processes have allowed to characterized framboidal, pentagonal, octahedral, tetrahedral, dodecahedral and icosahedral pyrite forms as originating from biogenic factors (Astafieva et al., 2005) This type of geometry encompasses all the euhedral morphologies described above for the Dead Sea core and mat. Framboidal pyrites are formed of these euhedral structures as
113 Chapter IV. Influence of microbes on the sedimentary record well as from spheroids of micrometric scales (Wilkin and Barnes, 1997), as observed in the core (Fig. 4.4B and 5D) and have been evidenced to be biomarkers in the sediment (Popa et al., 2004; Vuillemin et al., 2013a). Here, framboids are only observed very close to the gypsum crust within the microbial mat, while large euhedral pyrites dominate in the aragonite layers. Both microtextures have been shown to result from differential rates of pyritization (Wilkin and Barnes, 1996). In their demonstration of pyritization from Fe-monosulfides proceeding via Fe loss, the authors show that fast pyritization leads to framboidal pyrite while euhedrons would result in slow initial pyritization stage. Studies from Mediterranean sapropels show that framboids are associated to environments where HS- is produced by sulfate reduction (SR) within the sapropel, while euhedral pyrite form below the sapropel when HS- escapes from it before reacting with available iron. This is suggested to be caused by iron limitation (Passier et al., 1997) while high rates of sulfate reduction occur in the sapropel itself. Differential pyrite morphologies below the microbial mat may arise from similar processes. Intense sulfate reduction in the microbial mat could allow framboidal pyrite to develop. Excess HS- could diffuse away and react with iron monosulphide to form the observed large euhedrons in the aragonite lamina, at the interface with detritus laminae (Fig. 4.3). Iron liberation would be sufficient as derived from the neighbor detritus layer.
Fig. 4.7: Photographs of the cores (A) black layer exhibiting peaks in magnetic susceptibility and greigite occurrences near AAD intervals; (B) isolated S° concretions in reworked intervals and (C) S° concretions in disturbed and undisturbed AAD.
114 Chapter IV. Influence of microbes on the sedimentary record
In the core, very few framboids were observed as such (at 63 mblf only, see Fig A9 of the appendix). Fe-S minerals often occurred as micrometric individual spherules rarely aggregating into framboids. At given intervals, euhedral Fe-S similar (in morphology) to those observed in the mat, but often smaller in size could be observed. We conclude that sulfate reduction is occurring in the Dead Sea sediment, as witnessed by Fe-S minerals. Additionally, sulfur concretions were localized in the whole core (Neugebauer et al., 2014). General distribution rules cannot be defined, as is seems that they occur both in well preserved or seismically disturbed AAD, as well as in mass wasting deposits (Fig. 4.7 B-C). Different pathways could thus arise from their formation, from incomplete sulfate reduction as suggested by Bishop et al. (2013) or from oxidation of reduced sulfur species by turbidity events. They however support the role of microbial sulfate reduction in the accumulation of sulphide or S° in this sediment of the Dead Sea Basin. However, the fact that the chemistry of Fe-S rarely reach that of pyrite, and is generally close to greigite rises questions regarding the process of pyritization in this sedimentary environment. We address this issue in the next paragraphs.
4.6.2 Organic matter production and sulfate reduction
Communities inhabiting the gypsum sediments are mainly composed of highly adapted halophilic Archaea, relying mostly on fermentation of varied organic substrates (Thomas et al., 2014). On the opposite, communities inhabiting the aragonitic facies (AAD) seem to rely on the degradation of moderate halophiles biomass derived from the stratified water column (Ariztegui et al., 2015). EPS production could be related to a stress response of these moderate halophiles. It could also originate from sulfate reducing microbes potentially inhabiting these sediments, as they have been recognized to be important EPS producers (Bosak and Newman, 2005; Braissant et al., 2007). Degradation of these EPS by fermenters or sulfate reducing bacteria may release organic molecules such as aminated compounds (Mishra and Jha, 2009). They would be subsequently available for osmoprotectant synthesis or uptake by community members utilizing the low-salt in strategy (Roesser and Müller, 2001), or for other communities such as methanogens, as non-competitive substrate (Thomas et al., 2014), enabling more activity and organic matter degradation. The quality of organic matter is critical for microbial activity in deep lacustrine sediments (Glombitza et al., 2013; Ariztegui et al., 2015). In the Dead Sea, energy
115 Chapter IV. Influence of microbes on the sedimentary record requirements for osmotic adaptation select for metabolisms yielding the largest amount of ATP (Oren, 2001). In such competition, labile substrates and nutrient availability have a major impact. Water column communities and shallow sedimentary ones, like those observed in the microbial mat for example, will beneficiate from fresher and more labile organic matter and will subsequently be able to perform metabolic activity deeper ones cannot afford. The mat microcosm beneficiates both from the presence of a salinity gradient and from that of fresh organic matter. Among others, Dunaliella algae likely developed and survived in the pond after fresh water inputs to this small ephemeral pond (rain events occurred in December 2010; Fig. 4.2A and 2C). Mixing with Dead Sea water, dissolution of surrounding salt and evaporative process in such a shallow pond would quickly raise elemental concentrations to those observed at sampling time. The decay of these autotrophs provide large amount of highly labile organic matter to the shallow sedimentary community, allowing the development of the mat. Very little pyrite has been observed in the Dead Sea core. Although framboids and spherulites have been occasionally found, the majority of iron sulphides lie around the mackinawite and greigite chemistry (Fig. 4.6). Iron monosulphides have been acknowledged as important precursors towards the formation of pyrite (Berner, 1970; Canfield et al., 1992; Wilkin and Barnes, 1996). Pyritization has been observed to occur through different pathways : 1/the polysulfide pathway consisting in adding S° to the FeS precursor (Rickard, 1975; Luther, 1991) ; 2/the ferrous iron loss pathway (Wilkin and Barnes, 1996) where oxidants such as O2, nitrate, Fe(III), Mn(IV) or organic matter
2+ allow the release of a Fe ion, or 3/ the H2S pathway consisting in a solid state reaction (Drobner et al., 1990; Rickard and Luther, 1997; Pósfai et al., 1998). Transformation of mackinawite to greigite, and greigite to pyrite necessitate similar reactants, although kinetic limitations highlighted by Hunger & Benning (2007) prevent the full transformation of mackinawite to pyrite in S° limiting conditions. Our data suggest a transitional path from mackinawite to greigite to eventually pyrite in the Dead Sea core. Although no general rate can be observed as we do not see a correlation between pyrite content and depth, it seems that progressive sulfidation as suggested by Sweeney and Kaplan (1973) and Schoonen and Barnes (1991) allows pyrite formation in the Dead Sea. This sulfidation is most likely the result of microbial sulfate reduction in the Dead Sea sediment. Such process is acknowledged in the present Dead Sea (Häusler et al., 2014) where it can benefit from very sharp salinity gradients formed, for example by
116 Chapter IV. Influence of microbes on the sedimentary record
Fig. 4.8: Photographs of EPS morphologies in the mat (A) Cryo-SEM picture of EPS sheet in between gypsum crystals. Note the wrinkly cerebroid structure clearly defining the biofilm. (B) Cryo-SEM picture of an aragonite stellate cluster embedded in sheet-like EPS. (C) Regular SEM picture of sheet like EPS covering detrital sediment. Globules are clay minerals, needles are aragonite. (D) Regular SEM picture of sheet-like EPS, torn and folded as it can occur in the core, here at 23.59 mblf in core 1-A-13. underwater fresh water springs. Rates in the sediment are however very slow because of the high salinity and low fresh carbon inputs. It means that the production of sulphide from sulfate, which is allegedly infinite in most of the deep Dead Sea sediment (as observed from gypsum saturation), is limited. Recently, Bishop et al. (2013) suggested that sulfate reduction in the Dead Sea sediment does not even proceed all the way to
H2S, because of the lack of electron donor. Such hypothesis lies on the sulfur isotope composition of native sulfur concretions of the Lisan Formation. They argue that the absence of pyrite in the sulfate rich Dead Sea sediment is linked to this incomplete SR terminating at S°, thus allowing coexistence of mackinawite, and greigite and sometimes pyrite, as demonstrated by Hunger and Benning (2007) in limiting sulfur conditions. Our data are in line with such hypothesis. The presence of framboidal and euhedral
117 Chapter IV. Influence of microbes on the sedimentary record pyrite (Fig. 4.5A and 5C) in the microbial mat can be supported by complete SR allowed by the salinity gradient in the pond and fresh OM provided by Dunaliella algaea. In the core, we suggest that pyrite occurs only when enough fresh organic matter is transported to the sediment, with potentially sufficient dilution of the hypolimnion (as suggested by Lazar et al., 2014). Such events could have occurred at the end of the climax of glacial periods in the Dead Sea Basin. Pyrite could thus be used as a productivity proxy in the deep Dead Sea sediment. Differential occurrences of Fe-S minerals and in particular greigite in halite and AAD facies also suggest variations in the S and C cycles in both limnological settings. During meromictic periods, sulfate reduction is limited but outcompetes Fe reduction due to lowered porewater salinity and probably higher productivity in the diluted epilimnion. Greigite rims were observed around Ti- magnetite (Ebert, personal communication), the latter being the source of iron as marked by dissolution traces on its surface, similar to those observed in Lake Kinneret (Nowaczyk, 2011). In halite rich sediments, if detritus are absent, no Fe-S are observed as no Fe source is available. In the case of detrital inputs to the lake, Fe-S mineralizations are observed around gypsum (Kiro, personal communication). This suggests that sulfate reduction is here even more limited. No autotrophic production occurs during periods of halite precipitation. Sulfate reduction rates are lowered by the lack of production of fresh organic and a higher salinity. This only allows for the precipitation of Fe-S around the sulfate sources. Potentially, iron reduction out-competes sulfate reduction in this type of sediment.
4.6.3 Microbial effects on early diagenesis in the deep Dead Sea sediment
Precipitation of aragonite occurs in the Dead Sea when Mg-Ca brines of the hypolimnion encounter carbonate-loaded fresher water of the epilimnion. This is thought to occur either through the simple adding of these carbonates (Barkan et al., 2001), or through alkalinity rise during CO2 escape caused by evaporation, or by CO2 consumption by photosynthetic blooming organisms (Neev and Emery, 1967; Begin et al., 1974; Kolodny et al., 2005). Carbonate ions thus seem to be limiting the precipitation of aragonite in the Dead Sea. As a result, their production in the sediment or at the water sediment interface could be a potential trigger in the precipitation of aragonite, especially since associated metabolism promote alkalinity rise (Dupraz and Visscher, 2005). Dupraz and
118 Chapter IV. Influence of microbes on the sedimentary record
119 Chapter IV. Influence of microbes on the sedimentary record
Fig. 4.9: photographs of EPS structures in the mat and the core. (A) Cryo-SEM back- scattered electron photograph of an aragonite stellate cluster embedded in wrinkly EPS. Close up view (B) shows the tight relationship between aragonitic thin needles in the EPS matrix. (C) Nanoglobules on the surface of torn and folded EPS. In the back, EPS display vacuolar textures. (D) Aragonitic needles covered with EPS (as seen by torn and vacuolar textures) and with nanoglobule-like textures at their tips. Needles are thin, poorly defined and not smoothened here. (E-F) Aragonite stellate clusters embedded in EPS form core 1-A-2 at 2.74 mblf and 3.22 mblf. (G) Back-scattered electron imaging of Fe-S mineralizations embedded in EPS, in between aragonitic agglomerates (top) and gypsum mineral (bottom) from 2.74 mblf. (H) Aragonite stellate cluster at the surface of halite smooth surface at depth 337.62 mblf, in core 1-A-131. A-D from the mat, the rest from the core. All pictures taken in secondary electron imaging unless stated otherwise. coworkers (2009) have listed potential mechanism that would enhance calcium carbonate precipitation. Among them, complete sulfate reduction would permit the release of EPS-bound Ca2+ as well as carbonate ions by degradation of EPS, raising alkalinity and CaCO3 saturation index (Visscher et al., 1998; Visscher et al., 2000). Effects of sulfate reduction on alkalinity have been lately discussed by Meister (2013) and in particular the fact that the production of one H+ per mole of reduced sulfate also leads to pH decrease, hence to more carbonate dissolution than precipitation. Such model seems to be dependent on the type of organic matter used as electron donor (Gallagher et al., 2014). In the case of the Dead Sea however, degradation of EPS is not needed for releasing bonded calcium, as suggested in the process of biologically-induced mineralization (Dupraz et al., 2009), since calcium is already present in the porewater at extreme concentrations (Levy et al., in prep). The use of formate or hydrogen as electron donor for SRB could thus also promote carbonate precipitation under the form of aragonite (Gallagher et al., 2012), and is known to occur at high salinity for various species (Oren, 2010). Nanosphere presence (Fig. 4.9C), and what seems to resemble to transformation of the organic matrix to calcium carbonate mineral observed in the mat (Fig. 4.9D) may thus be influenced by the microbial production of carbonate ions in the sediment, from the available allochthonous or autochthonous organic matter. These nanoglobules have been recognized to be the initiating stage of calcium carbonate precipitation in numerous EPS-rich environments (Benzerara et al., 2006; Bontognali et al., 2008; Spadafora et al., 2010), or in association with sulfate reducing bacteria (Aloisi et al., 2006). They finally lead to the formation of stellate aragonite clusters, embedded in, or originating from, the biofilm sheet remains (Fig. 4.9D-F) as those observed in the marine stromatolites of Highborne Cay, Bahamas (Reid et al., 2000). The final aragonite
120 Chapter IV. Influence of microbes on the sedimentary record morphology is similar to other aragonite precipitates (needles or stellate clusters) in the Dead Sea. Clustering into stars may occur through EPS related nucleation, but also via nucleation on the surface of detrital minerals or bacterial surfaces (Spadafora et al., 2010). No direct morphological characteristic can thus be identified as the only result of the influence of EPS on aragonite precipitation. While potentially observable in the mat (but not recorded in the bulk aragonite carbon isotopes (Fig. A6), microbially influenced aragonite precipitation is relatively unlikely to occur in the core. First of all, no proof was observed for it. Absence of nanoglobules or of conclusive aragonite-EPS interactions and morphologies tends to discriminate against such pathway. Second, this pathway would depend on the complete organic matter degradation from microbial activity. Our data do not provide evidence for such pathway in the core. Sulfate reduction occurs at slow rates and/or seems to be incomplete, bringing limited effects on the alkalinity of the sedimentary micro-environment (Dupraz and Visscher, 2005). Its impact on the Dead Sea environment should be minor compared to the huge amount of aragonite precipitating in the water column: 15 mol.m-2.y-1 in Lake Lisan (Stein et al., 1997), and 1.4 mol.m-2.y-1 after heavy flooding of winter 1992 (Barkan et al., 2001). Stable isotopic measurements of bulk aragonitic carbon did not indicate specifically microbially induced fractionation, although some measured values are relatively low (Fig. 5.2 in chapter 5), which conducts us to minimize the impact of such pathway on the total aragonitic pool. However, they are important to better understand future diagenetic processes susceptible to affect porosity and OM degradation in hypersaline sediments. Greigite precipitation is diagenetic and will clearly influence the magnetic signature of the sedimentary record. This had already been highlighted by Ron et al. (2006) and (Frank et al., 2007). Greigite rims regularly observed around these minerals in AAD intervals, even in the early Holocene (Fig 7A) suggests the rapid reaction of soluble sulfide with dissolved iron originating from the detrital iron fraction. On the opposite, Fe-S precipitates have been observed in the halite sections of the core, only when gypsum is an associated evaporative phase (Kiro, personal communication). The detrital component seems to be the source of iron there, and Fe-S forms around the gypsum minerals. Different effects are thus observed, whether we observe sediments derived from arid or more humid periods. This early diagenesis is however erased by a second stage, if sediments ever outcrop and desiccate. No greigite but only pyrite has been
121 Chapter IV. Influence of microbes on the sedimentary record observed in the Masada sections of Lake Lisan (Bishop. et al, 2013) and halite intervals dissolve when outcropping. The iron and sulfur cycles are deeply influenced by microbes in the Dead Sea subsurface with phase changes from sulfate to gypsum to HS-, S0, and Fe-S mineralizations. The quality of the available organic carbon is critical to the formation of these phases. It also seems that the processes vary whether we are looking at halite rich intervals precipitating during arid periods, or AAD intervals characteristic of more humid episodes. Similarly to the effects on microbial communities (Ariztegui et al., 2015), freshwater inputs largely affect the precipitation and diagenetic processes in the deep Dead Sea Basin.
4.7 Conclusion
The Dead Sea is a unique system that can only to a certain extent be compared to other hypersaline systems. It possesses hypersaline system characteristics such as unique development of microbial communities, but at much slower rates, and with different limiting conditions (low sulphide, high Ca2+ and Mg2+ concentrations). The drilled record sheds light on the evolution of deep sediments before they undergo dehydration or dissolutive transformation. They imply diagenetic transformations occurring in the deeper part of the basin. The ICDP-core unique record hosts Fe-S mineralizations, emphasizing the existence of a microbial sulfur cycle influencing mineral precipitation and organic matter preservation. Fe-S minerals are relatively small and close to the greigite chemistry, supporting a slow metabolic rate for sulfate reduction (or an incomplete one) in the Dead Sea sediment. Comparison with an active microbial mat of the Dead Sea shore also allowed detecting the occurrence of exopolymeric substances in the sediment of the core. Based on morphologies mainly, they have the potential to serve as templates for aragonite precipitation at depth, indicating that aragonite may not only form in the water column conditions previously envisaged. Together, these findings support the existence of living microbes in the sediment of the Dead Sea, and remain the only proof for their metabolic activity. Finally, these results have strong implications on the fate of organic matter and on processes involved in mineralization and porosity development in the perspective of diagenetic evolution of hypersaline subsurface systems. They also support the microbial impact on the sedimentological and geochemical record of the deep Dead Sea sediment.
122 Chapter IV. Influence of microbes on the sedimentary record
Such observations must be taken into account in future paleoenvironmental reconstructions.
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126 Chapter IV. Influence of microbes on the sedimentary record
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127 Chapter IV. Influence of microbes on the sedimentary record
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Appendix Material
Fig. A6 Carbon isotopes of aragonite, detritus and aragonite mixed with detritus laminae for the microbial mat samples at En Qedem. Corresponding TOC is also presented in the 13 table with obtained ∂ CCaCO3
128 Chapter V. Influence of life on the Dead Sea geochemical record
“Pray for rain”
Heligoland, Massive Attack
129
130 5.1 General influence of the biosphere on the carbon cycle of the Dead Sea (and its precursor lakes).
5.1.1 Introduction to the Dead Sea carbon cycle
Being a terminal lake in an arid environment, the carbon cycle of the Dead Sea is largely driven by its evaporation/precipitation ratio (E/P). As discussed previously, fluctuations of lake level, forced by climate variations are caused by changes in the intensity of freshwater inputs and evaporative processes in the Levantine area, and even in the Middle East region (Bartov et al., 2003; Waldmann et al., 2009; Kushnir and Stein, 2010). These variations have reached several hundreds of meters during the last 200 ka (Waldmann et al., 2007), and are visible to the human eye today through the fast retreat of the shoreline. They inevitably form the prime cause of any variation in the major ion chemistry of the Dead Sea, making it harsh to assess the effect of biological activity through their direct measurement. The current intense dynamic of the Dead Sea limnology can nevertheless be used as an asset, as major changes have recently occurred in the water column, and have been monitored despite the difficulties its chemistry induces. In this way, observations of algal blooms during dilution events and the monitoring of their evolutions by Aharon Oren and co-workers have allowed to understand the key features relative to intense life development in the Dead Sea (Oren and Shilo, 1982a; Oren et al, 1995). Direct measurements on the water during the turnover period, in the winter of 1972 and 1992 have also unveiled the physics and chemistry involved in the fate of the epi- and hypolimnion, and its mixed counterpart (Luz et al., 1997; Barkan et al., 2001). These data are today prevalent to reconstruct and distinguish life dynamics in the Dead Sea sediments. During the second part of the 20th century, the Dead Sea lake level has decreased regularly, first due to generally arid conditions, and also to the intense use of the Jordan water, its main freshwater source, by Israel, Jordan and the Palestinian Authority. The evaporation rate is also intensified by the derivation of lake water to evaporation ponds in the Southern lake for salt extraction, by Dead Sea Works and Arab Potash Company (Tahal Group and Geological Survey of Israel, 2011). Whatever the key players are, the E/P ratio remains the impacted feature, from which derives other parameters such as salinity fluctuation, organic matter and nutrient inputs, setting or destruction of water column stratification. To understand the lake carbon cycle, let us first describe the main
131 inputs, outputs and sinks for C in the Dead Sea realm. Inputs of carbon in the Dead Sea are coming mainly from the Jordan and Mujib rivers (through DIC, DOC and POC), and punctually from « wadis » activated by heavy rainfalls over the Judean mountains and the Jordan plateau. They have eroded their way through canyons across the eastern and western shoulders of the Dead Sea Basin (e.g. Neugebauer et al., 2014). The presence of underwater springs along the eastern and western shores of the Dead Sea are also well documented and were thought to have a strong influence on the origin of allochthonous organic matter and carbon (Luz et al., 1997). Recent studies show that these springs may play a role in the global nutrient and carbon cycle of the Dead Sea and should be taken into account for further modeling (Ionescu et al., 2012).
5.1.2 Carbon isotopes in the Dead Sea realm
Being an endorheic lake, the only output for the Dead Sea water is through evaporation. It plays a key role in the abiotic carbon cycle. When E/P increases, the salinity of the surficial water increases too, which lowers CO2 solubility in water, subsequently enabling CO2 escape. This process of CO2 escape is described as the main contributor to alkalinity rise and has been invocated as the trigger for authigenic aragonite precipitation (Stein, 2001; Kolodny et al., 2005). Aragonite is the main carbon sink in the Dead Sea, with estimation of 1.4 mol.m2.yr of carbon sedimented after the 1994 flood, accounting for ca. 70 % of the total inorganic carbon input (Barkan et al., 2001). Other carbon sinks may be created by photosynthetic activity in the upper mixed layer. Dunalliela parva blooms are well documented in the Dead Sea and, according to experiments, occur when the epilimnion reaches a 70% dilution by freshwater in rainy winters, like that of 1992 (Oren et al., 1995; Oren, 1999). Carbon dioxide is fixed by the organisms, which also increase alkalinity and may induce aragonite precipitation. Two sinks of carbon may be created in that manner, through biomass formation and aragonite precipitation. The created biomass is either partially or completely remineralized by heterotrophic microorganisms such as those described in chapter II and III (Oren et al., 1995 ; Thomas et al., 2014) or may sink and be buried in the sediment. The exact extent to which all of these processes affect the global Dead Sea carbon cycle is hard to estimate and beyond the scope of this study. We mainly focus on the putative effect of biological activity within this realm, in order to assess if it can
132 overprint intense abiotic signatures, which could be used for paleoenvironmental proxies. In this purpose, carbon isotope analyses bring key information as they do not take into account the intense concentration fluctuations often observed in the Dead Sea, 13C between 12C and 13C isotopes (1). but only the ratio δ
= 13퐶 1 × 1000 (1) 12 � 퐶�푠푠푠푠푠푠 13 13퐶 훿 퐶 � 12 − � � 퐶�푉푉푉푉 Moreover, fractionation of carbon isotopes (2) is often larger when life takes part in phase changes.
= (2) 13 13 푏푏푏푏푏푏푏 퐴 푎�푎 퐵 푝ℎ푎푎� 퐵 푝ℎ푎푎� 퐴 This is the case for biological∝ fixation where훿 퐶 fractionation− 훿 퐶 indexes can reach -30 ‰ for Rubisco carboxylation (O’Leary, 1981) or for the reductive acetyl-CoA pathway domiantant in the Euryarchaeota branch (Berg et al., 2010). Remineralization of this 13C depleted organic carbon 13C values. This is compared to only -0.9
‰ for CO2 dissolution andgenerally +2.7 ‰ induces from carbonate low δ ions to aragonite (at 25 °C, Romanek et al., 1992). Hence, depending of equilibrium and initial conditions, carbon isotopes can help infer dominant pathways. By compiling data from carbon concentrations and isotopes monitored in the Dead Sea water column (Luz et al., 1997; Barkan et al., 2001), mass balance calculations during Dunaliella blooming periods (Oren et al., 1995) and lake level fluctuations (Gertman and Hecht, 2002), it is possible to approach the effect of microbial activity on water chemistry. Until winter of 1978, the Dead Sea was for a long period in a stratified regime. Under such conditions, the epilimnion was oxic and had a 13CDIC around 1 and 2 ‰ (Neev and
Emery, 1967 ; Fig. 5.1). It was relatively decoupledδ from the anoxic hypolimnion for 13CDIC lay below -7 ‰ (Nissenbaum, 1975). The study of Luz et al., (1997) whichfocuses δ on the changes of the isotopic ratios of dissolved carbon during the mixing and renewed stratification events that occurred in the Dead Sea between 1978 and 1985, and between 1992 and 1994. In early 1979, the Dead Sea water column completely ventilated (Steinhorn et al., 1979) and isotopic values of the DIC at the surface and below 200 m homogenized and dropped to ca –4 ‰, supposedly due to mixing with water depleted in 13C by sulfate reduction activity (Luz et al., 1997). The humid 1979-
133 1980 winter reestablished stratification until December 1982. A bloom of the red- pigmented green algae Dunaliella parva followed the dilution of the epilimnion in summer 1980 (Oren and Shilo, 1982) 13CDIC at the surface. This was followed by a rapidand causeddecrease a strongpotentially increase owing in the to δ subsequent oxidation of the produced biomass, either by oxygen or through the intense development of heterotrophic Halobacteria documented in the following months (Oren, 1983). After disappearance of the upper mixed layer (between 1983 and 1986), values
13CDIC remain relatively stable and positive in average. Small and short-scaled changesof δ are observed at both depths and are allegedly caused by seasonal stratification and mixing (Luz et al., 1997). Monitoring of the Dead Sea isotopic carbon resumes right after the renewed stratification of 1992. At this period, heavy rains allow a new Dunaliella bloom to develop, inducing again a strong 13C enrichment of the surface water DIC. Calculated fractionation caused by this event in a 7 m thick mixed layer on the whole lake surface is + 8.5 ‰ (Oren et al., 1995), leading to values relatively similar to what has been observed at the surface before the 1980 bloom. Depletion of 13C follows the blooming event, and is probably caused by Halobacteria response, CO2 escape to the atmosphere, vertical mixing and aragonite precipitation. The aforementioned abiotic factors are invoked for the following seasonal fluctuations in 1993 and 1994 (Barkan et al., 2001). Overall, carbon isotopic variations are induced by precipitation and evaporation, as outlined by the lake level curve on Fig. 5.1. They induce changes in lake stratification and quick biological response that lead to short- 13CDIC values. These extrema signatures could arguably be used as tracerstermed for extreme intense δbiotic activity in the Dead Sea epilimnion. However, they are only slowly transmitted to the bottom water, where they could be potentially preserved in the sediment pore water if not altered through diagenesis. Indeed bloom signatures seem to only occur in a decoupled epilimnion and the hypolimnion evolves individually in such settings, before turbulent mixing and density gradient allows transfer of this 13C depleted water mass to the bottom (Lazar et al., 2014). Hence, such geologically ephemeral signatures are not likely to be transmitted to the larger hypolimnion carbon pool. They could however be recorded in the sediment by precipitation of aragonite from the epilimnion.
134 5.1.3 Aragonite precipitation 13C signature in the core
In the Dead Sea, aragonite is andthought its δ to form with Ca2+ originating from the Ca-Mg-rich
Dead Sea brine, and CO32- brought with incoming freshwater. The timing of this aragonite precipitation is however controversial. Work based on carbon fluxes modelling concluded that in the case of the modern Dead Sea, aragonite precipitation should occur directly upon mixing of brine and freshwater (Barkan et al., 2001). This is different from the lake Lisan period, for which it is largely accepted that aragonite is controlled by evaporation and thus is more likely to occur in summer (Neev and Emery,
1967; Begin et al., 1974). Kolodny et al. (2005) stress the fact that CaCO3 precipitation may not only be caused by CO2 outgassing. Phytoplankton blooms, may also remove CO2 through photosynthetic activity, and thus raise alkalinity, subsequently promoting carbonate precipitation. Recent blooming events show that this process should not be neglected. The absence of aragonite minerals in the lake in May 1992 however supports the idea of a winter precipitation (Luz et al., 1997), and could minimize the impact of blooms on short aragonite precipitation events in the modern Dead Sea. Prolonged stratification episodes allowing planktonic development would however still affect the 13C of aragonite. In both cases, precipitation of aragonite should occur in the epilimnion,δ where carbonate ions are available, and would reflect processes of carbon fractionation at stake in this layer. Although not available from the DSDDP cores yet, extensive carbon and oxygen measurements together with other isotopic studies from aragonitic deposits have been realized from outcrops in the Masada area and have allowed reconstructions of lakes Lisan and Samra environments (Kolodny et al., 2005; Waldmann et al., 2007). A long- term (ca. 55 ka) 13C of aragonite across the Lisan Formation has also been tentatively increaseattributed in theby δ Kolodny et al. (2005) to biological activity by photosynthetic organisms. They are hypothesized to be soft-bodied algae similar to Dunaliella as no fossils were recovered from the sediment. These data tend do show that life can leave their imprint on the carbon cycle of the Dead Sea and that aragonite intervals in particular should be looked carefully before undergoing any paleoenvironmental study. In order to test this hypothesis, AAD intervals were selected within the available core- catcher sediment sampled during the drilling campaign. Aragonite minerals from white AAD laminae were carefully sampled. Several laminae from an identical interval were
135 taken in order to avoid unrepresentative values. They were then rinsed three times with ultrapure water and dried. Carbon and oxygen isotopes were measured on a Thermo Finnigan Delta Plus XL at the stable isotope laboratory of the Institute of Terrestrial Surface Dynamics, University of Lausanne, Switzerland. Results are shown in per mil VPDB, and error margin is ±0.05 ‰. Carbon isotopes are generally negative, with
Fig 13C variations of Dissolved Inorganic Carbon (DIC) at 2 and 200 m below lake level (m bll) and interpreted causes, between summer 1963 and summer 1994. Data compiled. 5.1: ∂ from Luz et al. (1997), Oren et al. (1995) and Barkan et al. (2001). Lake level reconstruction from Gertman & Hecht (2002).
136 average at -2.05 ‰ (Fig. 5.2). The maximum value reaches 3.51 ‰ at the bottom of the core, while minima are reached at depths of nearly 200 m below lake floor, with around -7.4 ‰. Minimum values are also found at depths of 58, 65, and 147 m blf (ca. -7 ‰). No trend is clearly observable though, and these results can hardly be interpreted through the curve shape since the scatter of data points along depth is too large to mean anything. Additionally, quick changes described before can lead to such scatter of points. Continuity is subsequently needed to constrain the carbon isotopic variation of aragonite laminae. A carbon isotope dataset of higher resolution will be available within the scope of DSDDP publications. What we will mostly focus on here is the minimum values, which could be linked to fractionation observed in the lake surface during periods of stratification (after photosynthetic activity), or after active degradation of organic matter in anaerobic conditions. Given the isotopic signature of freshwater from the Jordan and from floods (~ -7.7 ‰ as taken from Barkan et al., 2001) and the fractionation index from DIC to aragonite (2.7 ‰ at 27°C ; Romanek et al., 1992), minima values for aragonite are hardly reachable without additional recycling of carbon by microbial agents, if equilibrium is assumed. This proxy disturbance will be further discussed in the following chapter, which addresses the specific interval between 50 and 90 m in the core. However, it is here confirmed that microbial degradation of organic matter in the water column can influence the 13C of aragonite and disturb this proxy when used solely for climatic reconstructions.δ
13 δ Caragonite (%o) -8 -6 -4 -2 0 2 4 6 0
50
100
150
200
250
300 Fig. 5.2: Carbon and oxygen isotopes 350 from aragonite taken from AAD core- 400 catcher intervals, for core 1A.
450 Depth (m blf) 500 137 5.2 Anaerobic oxidation of methane disturbed organic proxies in the Early Holocene Dead Sea
This chapter is in preparation for submission in a peer-reviewed international journal as Thomas C., Levy E., Antler G., Neugebauer I., Grossi V., Sivan O., Yechieli Y., Gavrieli I., Turchyn A., Stein M., Brauer A., Ariztegui D. and the DSDDP Scientific Team ; Anaerobic oxidation of methane disturbed organic proxies in the Early Holocene Dead Sea.
Abstract
The Dead Sea Deep Drilling Project is an ICDP sponsored project that aims to reconstruct the paleoenvironments of the Dead Sea Basin. A study of multiple proxies derived from the pore water of drilled sediment, from sedimentological description and from lipid biomarkers highlights enhanced biological activity in the sediment dated from the beginning of the Holocene. Concordant release of light carbon isotopes and preferential mineralization of light sulfur and oxygen isotopes indicate a biological signature in the early Holocene, during a relatively humid period. Organomineralization under the form of euhedral pyrite morphologies suggest an increase sulfate reduction activity in one specific interval. This interval comprises a set of lipidic biomarkers indicative of anaerobic oxidation of methane. Among others, the presence of hydroxyarchaeol, pentamethyleicosene and extended archaeol highlights the reworking of biogenic methane by potential ANME Archaea. The combination of intense lake level drops and
Fig. 5.3 : Location of the Dead Sea and DSDDP drill sites
138 renewed stable meromictic conditions seem to provide the setting for the recording of microbial influence on organic proxies, and subsequently calls for enhanced attention towards biological processes when using hypersaline environments for paleoclimatic reconstructions.
5.2.1 Introduction
In the framework of the ICDP-sponsored Dead Sea Deep Drilling Project, a paleoenvironmental study is conducted through a record of several cores retrieved during winter 2010-2011 in the Dead Sea Basin. Around 450 m of sedimentary cores encompass both humid aragonitic rich glacial periods and arid halite dominated interglacial periods (Neugebauer et al., 2014). It will allow reconstructing climates of the last ca. 220 ka at a very high resolution. However, recent studies in lake subsurfaces have revealed the importance of considering biological effects on paleo-environmental proxies before undertaking paleoclimatic reconstructions (Ariztegui et al., 2015; Vuillemin et al., 2013; Chapter IV). Mainly through its name, the Dead Sea has regularly been considered as void of life, and influence of biological activity has often been overlooked. Attempts at addressing such issues have occurred (Nissenbaum, 1975; Luz et al., 1997; Kolodny et al., 2005), and have been followed by detailed investigations of its biosphere. Work by Aharon Oren, who followed the pioneer steps of Benjamin Elazari Volcani, have established the conditions for life development in the modern Dead Sea (Oren et al., 1995) and described its prime inhabitants (e.g. Oren and Shilo, 1982; Oren, 1983; Oren, 1993). Attentions through quickly developing microbial ecology techniques have allowed to address diversity and survival in the Dead Sea water (Bodaker et al., 2010; Rhodes et al., 2012), at its interface with the sediment (Ionescu et al., 2012) and deep in the sediment (Thomas et al., 2014), enlightening the sustained presence of microbes in such extreme environments. In this way, the putative influence of microbes should not be neglected when using proxies that could be disturbed by biological activity. Torfstein and co-workers have in the past highlighted such effect. Sulfur isotopes in disseminated gypsum of the Lisan formation, in the Dead Sea Basin, were indeed evidenced to bear the imprint of sulfate reducing activity (Torfstein et al., 2005; Torfstein et al., 2008). Particular attention has thus arisen towards the carbon and sulfur
139 cycling of the Dead Sea subsurface, as genes potentially linking to sulfate reduction, fermentation and methanogenesis have been evidenced in the sediment (Thomas et al., 2014). By using a combination of isotopic and organic geochemistry data, we intend to provide evidence for life influence in the Dead Sea sedimentary record, and bring light to a particularly intense event of the early Holocene in the Levantine region.
5.2.2 Geological setting
The Dead Sea basin is located in the Levantine region (Fig. 5.3), and was formed tectonically along the transform boundary between the Arabian and Sinai plate, through a pull-apart structure within the Jordan rift valley (Garfunkel and Ben-Avraham, 1996 and references therein). The Dead Sea is a terminal lake within this basin, lying at 427 m below sea level (2013) and constantly retreating since several decades (Abu Ghazleh et al., 2009). Its only permanent tributaries are the Jordan River and the Mujib River, which carry most of the freshwater inputs to the Dead Sea (Greenbaum et al., 2006). Heavy rainfalls over the Judean Mountains and Jordanian Plateau activate ephemeral wadis transporting additional water through flashfloods. Today, the precipitation-evaporation ratio (P/E) is largely negative resulting in shoreline decrease of a meter per year, and constant halite precipitation from the water column. Concordantly, halite-rich facies are dominant in interglacial periods, often mixed with detrital marls (Neugebauer et al., 2014). In the past, periods of higher lake stand have allowed the stratification of the lake with dilution of its upper mass by freshwater. Aragonite was then the main evaporitic phase, forming a varve-like alternation with detrital material brought to the lake during winters (Stein et al., 1997). This facies (AAD) largely dominates in glacial periods. During periods of decrease in lake water level, precipitation of gypsum occurs as a transitional phase between aragonite and evaporite (Torfstein et al., 2008). Massive gypsum deposits (known as the Upper Gypsum Unit in the outcrops) notably marks the end of the Pleistocene that was characterized by an abrupt drop in lake level (Torfstein et al., 2013).
5.2.3 Material
The sediment used for measurements originates exclusively from the core 5017-1A, retrieved from the middle of the Dead Sea at 300 m water depth (Fig. 5.3). Pore fluids were extracted from core catchers on the day of drilling as described by Lazar et al.
140 (2014) 13CDIC. Samples from opened cores during the core-opening parties in June and Octoberfor δ 2011 at the ICDP facilities in GFZ Potsdam (Germany) were squeezed with a hydraulic press pore water extraction system (2-column bench top laboratory press, 22 13C tonof aragonite max load, were Carver all Inc., sampled USA). duringSamples the for drilling lipid extraction, campaign, SEM after investigation each day of and drilling, δ using sterile tools, and kept in the freezer until further processing. Main characteristics of samples used for lipid biomarkers are given in Table 5.1.
5.2.4. Methods
Geological description
Detailed lithological description was performed using high-resolution scanned images of the core halves, µXRF profiles and microscopy using smear slides. Scanning Electron Microscope investigation was done on few samples after regular drying on a Jeol®144 JSM-7001 FA at the University of Geneva. Samples were mounted on an aluminum stub with double-sided conductive carbon tape. An ultra-thin coating (15 nm) of gold was applied by low vacuum sputter coating. Energy Dispersive X-ray spectroscopy and back- scattered electron method were used for Fe-S mineral detection. High resolution magnetic susceptibility was acquired on freshly opened and cut core halves at the GFZ in Potsdam, Germany (see Neugebauer et al., 2014 for more details).
Isotopic measurements on porewater
Subsamples
Allfor extractedDIC carbon pore isotope water analyses fluid was were filtered immediately through transferreda 0.45 μm syringe into 20 -filter.mL pre -poisoned airtight syringes (HgCl2 powder) to terminate bacterial activity. About 0.2 mL of each groundwater sample was transferred into a He-flushed vial containing H3PO4 for the headspace measurements of 13CDIC by conventional isotopic ratio mass spectrometer
(IRMS, DeltaV Advantage; Thermo,δ Waltham, MA, USA) with a precision of ±0.1‰. Subsamples for sulfate isotopes subsamples was acidified and purged with argon gas to
34Ssulfate 18Osulfate. Extracted pore removewater samples any sulfides had sulfate prior toprecipitated being analyzed as barium for δ sulfate (barite)and δ by adding a saturated barium chloride solution. The barite was subsequently rinsed with acid and deionized
141 water and set to dry in a 50 °C oven. The sulfur and oxygen isotope composition of the pore water sulfate were analyzed in the Godwin Laboratory at the University of Cambridge. The barite precipitate was pyrolyzed at 1450 °C in a Temperature Conversion Element Analyzer (TC/EA), and the resulting carbon monoxide (CO) was measured by continuous flow GS-IRMS (Delta V Plus) for its 18Osulfate. For the 34Ssulfate analysis the barite was combusted at 1030 °C in a Flash Elementδ Analyzer (EA),δ and resulting sulfur dioxide (SO2) was measured by continuous flow GS-IRMS (Thermo, Delta
V Plus). Samples for 18OSO4 were run in replicate, and the standard deviation of these
18 replicate analyses wasδ used as the external reported error (~0.5 ‰). The Osulfate values are reported vs. VSMOW and corrected for two barite standards ofδ known 18Osulfate that were run at the beginning and end of each set of samples (NBS 127
18 18 34 δ Osulfate = 8.6‰ and EM barite Osulfate = 15‰). Ssulfate results are reported vs. the δCanyon Diablo Troilite (VCDT),δ and the error wasδ determined using the standard deviation of the standard (NBS 127) at the beginning and the end of each run (~0.4‰).
The 34SSO4 was also corrected to two standards of known sulfur isotope composition,
NBS δ127 (20.3‰) and EM barite (12‰).
Lipid analysis
Samples were freeze-dried, ground and extracted through sonication cycles (methanol twice, methanol/dichloromethane (1:1) twice and dichloromethane three times). Sulfur was removed by activated copper. Precipitates were filtered out and lipids separated over an inactivated silica column into five fractions of increasing polarity. Fraction F1 was eluted with hexane/dichloromethane (9:1), fraction F2 with hexane/dichloromethane (1:1), fraction F3 with dichloromethane, fraction F4 with ethyl acetate and fraction F5 with methanol. Fractions F3 and F4 were silylated with pyridine/BSTFA 1:1 (v/v). Fraction F5 was trans-esterified as follow: fraction was incubated at 60°C overnight with toluene and H2S in methanol (2%). NaCl 5% and hexane:dichloromethane were then added and the organic upper phase extracted three times, washed with NaHCO3 (2%) and dried with sulfate. It was then silylated with pyridine/BSTFA 2:1 (v/v). Mass spectra were obtained by gas chromatography-mass spectrometry (GC-MS) using a MD800 Voyager spectrometer interfaced to an HP6890 gas chromatograph equipped with an on-column programmed at with the following parameters: injection at 60 °C, 30 sec equilibration, 20 °C /min to 130 °C, then 5 °C /min
142 to 250 °C and 3 °C/min to 300 °C. Identification of compounds was based on various gas chromatogram retention times and comparison of mass spectra, with known compounds and published data
5.2.5 Results
Lithology, magnetic and geochemical profiles
The description of the core has been detailed from that of Neugebauer et al. (2014) through observations of high resolution images and µXRF scanning of the core halves. Based on changes of dominant facies and chemistry, intervals (i) of changing precipitation over evaporation ratio (P/E) have been defined. During dominance of aragonitic facies with increasing sulfate concentrations, P/E are interpreted as positive, while halite/gypsum dominated facies with decreasing sulfate have been defined as negative P/E periods. Fluctuations of magnetic susceptibility are also used in this process (see Neugebauer et al., 2014 for more precision). These periods are defined starting from the massive gypsum unit at ca 100 m (Fig. 5.4). Before 100 m, AAD facies dominated, being either greenish or intercalated with detrital layers, typical of the glacial period and lake high stand. Slightly above 100 m, gypsum becomes the dominant
sample depth facies salinity TOC Ca2+ Mg2+ Cl- Na+ (m) (% TDS) (%) (mmol/ (mmol/ (mmol/ (mmol/ KgH2O) KgH2O) KgH2O) KgH2O) Surface 0.2 hh 33.6 0.02 438 2038 7050 1817 S17 64.26 AAD 32.2 0.67 184 784 6400 4336 S18 67.09 ld 31.4 0.48 110 727 6173 4356 S19 76.76 hd 33.2 0.60 271 1310 6745 3326 S21 91.04 gypsum 24.9 0.78 74 321 5438 1795
Table 5.1: principal characteristics of samples used for lipid biomarkers. Surface sediment originates from the surface of the DSDDP core 5017-1-A. Facies given as described by Neugebauer et al. (2014). Major ion concentrations from Lazar et al., 2014 and Levy et al., (in prep. TOC taken from chapter II. For S19, sufficient pore water could not be extracted and data are subsequently given from a sample located 60 cm below S19 : S19 at 76.76 m blf, major ion concentrations (in italic) measured at 76.16 m blf.
143 fraction. During this interval, sulfate values drop very quickly, as SO42- is precipitated through CaSO4-2H2O (i1). The concentration rises again while aragonite precipitation resumes together with packages of detrital sediment right above 90 m (i2). Negative peaking of 13C is
34 observed within this interval, and concordant with small increase in S.훿 Halite precipitation takes over (lh/hh) between 87.5 and 80 m, defining a second period훿 where evaporation dominates (i3). Occurrence of a 1.5 m-thick deposit of disturbed greenish AAD then reveals a strong input of freshwater, which is not correlated with peaking in sulfate as resolution is too low in this interval (i4). This event seems intense and relatively short as halite dominance resumes until 74 m (i5). Large detritus dominates aragonitic mud (ld), together with a steady increase in sulfate concentration until 62 m,
Fig. 5.4 : Lithological and geochemical profile along the 120 first meters of core 5017-1A. Periods i1 to i6 have been defined by a combination of lithological and geochemical observations. The lithological description and ages detailed from Neugebauer et al. (2014). Water balance is a result of the estimated ratio between P precipitation and E evaporation. Isotopic signatures are influenced by variations in freshwater inputs, evaporation and microbial activity (Sulfate reduction mainly), shifting them away from the general Dead Sea brine signature (in between dashed lines for 34Ssulfate. Depth unit is in meters below lake floor, and magnetic susceptibility is 10-6 SI plotted on a logarithmic scale. Gaps in the lithological description show absence휹 of core recovery during drilling. The latter is often caused by occurrence of hard halite deposits that are harder to drill (Neugebauer et al., 2014). Error bars are smaller than symbols.
144 implying renewed dominance of precipitation (i6). During this interval, maximum values 34 18O of sulfate, respectively. ofCarbon 40.21 isotopes ‰ and 20.44 also fluctuate ‰ are attained greatly for before ∂ S andattaining ∂ a minimum of -15.63 ‰ at the top of the interval. Magnetic susceptibility also stabilizes at relatively high values. Above this, magnetic susceptibility drops and sulfate decreased again in a large interval where occurrence of detritus mixed with halite (hd) and common gypsum appearance in single laminae indicated renewed aridity (i7). Above 50 m, sulfate increased again and aragonite mixed with detritus mud (ld) became dominant (i8).
Lipids
Four samples have been selected from different lithologies and intervals have been analyzed for their lipid composition (Table 5.1). Two of them have been taken from the i6 interval (S17 and S18). S19 is taken from a mixed halite and detritus interval in the i5 interval, and S21 from a gypsum unit interpreted as the corresponding interval for the capping “Upper Gypsum unit” marking termination of the Lisan formation (Lazar et al., 2014 ; Neugebauer et al., 2014). They show few differences in general, except for sample S21. For all samples, the hydrocarbon fractions F1 are dominated by alkane profiles, with general odd-over-even number dominance, increased in sample S18, and always centered on C29 and C31 n-alkane (Fig. 5.5A). Phytane is observed on sample S17 and S18 only, as well as occurrence of C17 n-alkane. Fractions F2 are very similar for S17 and S18, with elution of part of F1 observed through occurrence of n-alkanes (Fig. 5.5B). Major features are the detection of irregular isoprenoid compound interpreted as pentamethylicosene with two double bonds (Pmi) in sample S17 only (Fig. A7 in appendix). Additionally, C30 isoprenoid squalene and sterene are detected in S17 and S18. S19 is characterized by poor separation between the fractions F1 and F2. Typical alkenone profiles are additionally observed exclusively for this sample. Sample S21 displays wax esters characteristics of this sample. Fraction F3 shows linear alcohol profiles with even over odd dominance. Sample S17 and S19 are marked by important peaks of tetrahymanol, which remain relatively minor in S18 and S21 (Fig. A8). Fractions F4 are also generally similar from a sample to another. Main features are the occurrence of archaeol and extended archaeol (Ar and ext-Ar respectively) for all samples (Fig. 5.5C). The major difference lies in an increased peak at t = 61 min in sample S17, interpreted as silylated sn-2-hydroxyarchaeol (Ar-OH) through its mass
145 spectrum (Fig. 5.5D). Ratios of Ar-OH/Ar give respectively 0.03 and 0.02 for S17 and S18. Additionally, small peaks of compounds (a-g) interpreted as non-isoprenoidal macrocyclic di-ethers (mGD) with different number of methyl branches (Fig. 5.6) have been identified through their characteristic major fragmentation of m/z 145 (Fig. A9). Macrocyclic archaeol (h) has also been identified in S17 (Fig. 5.6).
SEM observations
Samples S17 to S21 have been observed under the SEM for the search of microbial traces. Keys for the recognition of such traces in the Dead Sea environments have been
Fig. 5.5: Lipid profiles of samples S17, S18, S19 and S21. (A) selective m/z=57 chromatograms of F1 (alkane). Note odd over even carbon number dominance of n- alkane, and high n=17 alkane peak for S17. (B) Partial total ion chromatograms of F2, with occurrence of Pmi for S17, alkenone for S19 and wax esters for S21. Note poor separation for samples S17, S18 and S21 with elution of n-alkane. (C) Partial total ion chromatogram of polar lipids. (D) mass spectrum of silylated Ar-OH and its non-silylated structure. -alkane, a-g: non-isoprenoid macrocyclic glycerol diether, Ar: archaeol, Ar-OH: sn-2-hydroxyarchaeol, ext-Ar: extended archaeol, h: macrocyclic archaeol, Ph: phytane, Pmi:(•) npentamethylicosene, Poll: pollution, Sq: Squalane, Sq*: squalene, Ste: Sterene.
146 taken from Part IV of this manuscript. Iron mineralizations have been obtained from all samples, generally as small size-minerals covered with precipitates, likely of evaporitic origin. Most iron-sulphide minerals are sub-micrometric spherules, which cannot be attributed to specific microbial or biological origin based on their shape only. Only sample S17 displays euhedral Fe-S minerals, sometimes in octahedron (fig. 5.7A), sometimes presenting geometric overgrowth (fig. 5.7A and 5.7B), diagnostic of diagenetic precipitation resulting in microbial sulfate reduction (see chapter IV for more details). Greigite and framboidal pyrite have also been retrieved at 69 m (Fig. A10 and A10). No EPS has been found in any of the samples.
5.2.6. Discussion 5.2.6.1 Decoupling of isotopes with the precipitation-evaporation ratio in i6
The use of the various evaporitic facies, together with sulfate concentrations allows a good view of relative lake level variations as described by Stein et al. (1997, 2001) and Neugebauer et al. (2014). Variations observed in carbon isotopes direct towards similar interpretations. During precipitation-dominated periods, the signature of freshwater mixing with the upper limnion water and that entering the hypolimnion through turbulent mixing (Lazar et al., 2014) lowers 13C values. Periods of intense evaporation, characterized by massive gypsum and halite∂ precipitation conversely correlate with 13C through CO2 escape triggered by salinity rise (Kolodny et al., 2005).
Whileincreases freshwater of ∂ input given by carbon isotopes signature is concordant with 34S of pore water sulfate actually increases. The precipitationlatter does not dominated support periods, a mixing the with∂ the normally isotopically light freshwater (Torfstein et al., 2008). These increases are however not regularly observed and occur
Fig. 5.6 : Proposed structures of macrocyclic glycerol diether compounds( a-g) and macrocyclic archaeol (h)
147
Fig. 5.7: large euhedral Fe-S morphologies obtained from sample S17 (white arrows) and interpreted as organomineralization. Note diatom fragments (black arrows) in photograph B.
34S = ca. 8 ‰) and i6. In the latter, it reaches 34S =
+40.21‰.only for i2 All (∆ values above 100 m below lake floor otherwise fall withina maximum the brine of ∂values defined for the Holocene Dead Sea (Gavrieli et al., 2001; Avrahamov et al., 2014). We thus conclude that other processes than the evaporation-precipitation ratio affect sulfur isotopes 18O of sulfate, which are here well correlated
34 .S. Similar Enriched conclusions sulfur isotopesarise from signatures ∂ tend to demonstrate a preferential depletionwith ∂ in light isotopes from the pore water. Bacterial sulfate reduction is commonly suggested in such situations and has been evidenced in the Dead Sea realm, through similar isotopic signatures measured from the massive gypsum layer (Torfstein et al., 2005; Torfstein et al., 2008). Here, values reach 20.75 ‰ in i2, and ca. 40 ‰ in i6, which is well above the maximum values previously measured in the Dead Sea sulfate fractions. A maximum is simultaneously reached in i6 by 18Osulfate suggesting a similar cause. Variations in lake level, with intense climatic fluctuati∂ ons has already been highlighted in the past through the relative variation of sedimentary facies, shoreline elevation and sulfur isotopes at the end of the Pleistocene-early Holocene (Stein, 2001; Bartov et al., 2002; Torfstein et al., 2008). In i6, pore water isotopes reach extrema never attained in stable periods of lake high stand for example, where more freshwater is flowing in the lake. A specific event, not directly correlated with the P/E ratio, has thus occurred during i6.
148 5.2.6.2 Information carried by biomarkers in the Dead Sea framework
Given the different lithological nature of the sediment and their relative occurrence within the core, biomarker profiles highlight features related to the sedimentary facies and limnology inherent to the Dead Sea. First, the similarity displayed by n-alkane profiles, with centering on C29 n-alkane suggests a general dominance of allochthonous organic matter, dominated by plant waxes (Meyers and Ishiwatari, 1993), whether lithology is of aragonite mixed with detritus, halite and detritus or gypsum with no apparent detritus. In the Dead Sea realm, organic matter is thus first controlled by the input of terrestrial organic matter. This has been highlighted by previous lipid studies in the Dead Sea Basin (Oldenburg et al., 2000). Allochthonous inputs easily hide autochthonous signatures given the low number of organisms generally expected in the Dead Sea realm. Other biomarkers that are equally distributed in all samples are those of halophiles and extreme halophiles. Ar and ext. Ar (Fig. 5.5C). They testify of the presence of such communities (Dawson et al., 2012) in the Dead Sea water column at time of sedimentation, or in the sediment. Members of the archaeal class Halobacteria, identified as the current sole inhabitants of the gypsum sediment S21 (Thomas et al., in review.; Thomas et al., 2014) are possible source organisms for these lipids. Wax esters, uniquely highlighted in sample S21, emphasize the harshness of the environment related to gypsum precipitation. Wax esters are generally accumulated by Bacteria in situations of high salinity, desiccation (Finkelstein et al., 2010) and nutrient starvation such as nitrogen starvation (Silva et al., 2007). The latter is common in sedimentary environments, and especially in the Dead Sea realm (Stiller and Nissenbaum, 1999). Interestingly, preservation of such esters is generally considered to be poor in sedimentary settings (Rontani et al., 1999), suggesting a possible in situ production from active communities of the gypsum sample (CT3 in Chapter II, and GY in Chapter III). Sample S21 has been taken at the top of a thick massive gypsum interval known to represent the abrupt transition (i1) between the relatively humid last glacial period, and the present interglacial period (Torfstein et al., 2008). This event, correlated to the “upper gypsum unit” of the onland Lisan Formation, has his upper limit dated at 14,103 ka ±190 years by radiocarbon (Neugebauer et al., 2014) and thus represents the termination of MIS2 (Torfstein et al., 2013). This very abrupt and arid period increased salinity and nutrient starvation in the lake through lack of freshwater inputs. It potentially resulted in the formation of wax ester as inclusion bodies in Bacteria
149 surviving in the lake or sediment. Occurrence of alkenones in sample S19 is surprising. This is to our knowledge the first retrieval of haptophyte markers in the Dead Sea realm. Dunaliella is the only eukaryote able to develop in the modern Dead Sea water (Oren, 2005). However, record of diatom fragments in sediment of the last glacial periods is common (Begin et al., 2004; Thomas et al., 2014) and indicates relatively fresher water episodes. In hypersaline environments, alkenone allegedly produced by halophilic algae related to Chrysotila lamellosa have been identified in saltern microbial mats (Rontani & Volkman, 2005). We thus hypothesize than in similar situations, haptophyte could have developed in the Dead Sea. Analogous conditions may have arisen during i4, where greenish AAD similar to those found during the Lake Lisan period were deposited. At this period, very intense freshwater input in the lake could have allowed algae to develop. The alkenone biomarkers of sample S19 may derive from communities that subsisted in the lake after this period, potentially in dormant stages or as dead cells Simulatenous recovery of tetrahymanol in this sample supports the preservation of signatures from communities related to fresher water input and hypolimnion anoxia (Sinninghe Damsté et al., 1995). Transport of these markers from slope instability may also be a potential reason as the highly disturbed AAD unit at i4 may also be a large chunk of glacial Lisan-related deposit that has been transported to the depocenter by mass movement. Differences pertaining to sample S17 and S18 only are of bigger interest to us as they potentially relate to the specificity of period i6. The occurrence of C17 saturated alkanes is suggestive of cyanobacteria or phytoplankton presence (Sachse et al., 2006) in the lake at time of sedimentation. It supports a stratification of the lake at that time, with a relatively fresh epilimnion and an anoxic hypolimnion, as suggested by the recovery of tetrahymanol in S17. Pentamethylicosene (PMI) is found in fraction 2 of S17 and S18. It is characteristic of methanogenic archaeal species Methanolobus bombayensis and Methanosarcina mazei (Schouten et al., 1997). It is also commonly accepted as a marker of anaerobic oxidation of methane (AOM 13C values
(Elvert et al., 2001). More recently, PMI has) wherebeen identified it shows as highly a specific depleted methanotrophic δ Archaea (members of the ANME group) derived biomarker (Niemann and Elvert, 2008; Chevalier et al., 2014). The enhanced recovery of hydroxyarchaeol in S17 could also be indicative of the presence of ANME (Hinrichs et al. (2000) as potential biomarkers of anaerobic methanotrophy. Such lipids can derive from methanogenic (Koga et al., 1998;
150 Bradley et al., 2009) and methanotrophic archaea (Rossel et al., 2008). Compound specific isotopes generally allow deciphering dominating source metabolisms as AOM related lipids display strong 13C depletion when using biogenic methane. The specificity of these samples is indicative of archaeal cycling of methane, being either methanogenic and/or methanotrophic. Methanogenesis has already been highlighted in the Dead Sea through metagenomics study of an AAD facies sample (Thomas et al., 2014) and is suggested to be a key element of this type of sediment throughout the sedimentary history of the Dead Sea Basin (see Chapter II of this manuscript). Upcoming compound specific isotopes could allow further characterization.
5.2.6.3 Anaerobic oxidation of methane
Combination of archaeol, hydroxyarchaeol and pentamethylicosene markers have often been described as classic signatures of methane seeps (Pancost and Sinninghe Damsté, 2003; Knittel and Boetius, 2009). Additionally, non-isoprenoid macrocyclic glycerol diethers are systematically found in environments rich in reduced sulfur, suggesting an origin from Bacteria involved in the sulfur cycle (Stadnitskaia et al., 2003; Pancost et al., 2006; van Dongen et al., 2007; Baudrand et al., 2010). The finding of this set of biomarkers points towards an association of methanotrophic Archaea (ANME) and sulfate-reducing Bacteria to oxidize methane by using sulfate as a terminal electron acceptor. The process of anaerobic methane oxidation works as follow:
CH4 + SO42- HCO3- + HS- + H2O (1)
Sulfate reducing activity would preferentially reduce light sulfur sulfate, leading to an 34 34 18O of sulfate in this enrichedinterval indicate∂ S in thes a residual similar poreprocess water. influencing Covariance these of ∂ Sisotopes. and ∂ Bacterial sulfate reduction (BSR) has been shown to influence in such way oxygen and sulfur from residual sulfates (Brunner et al., 2005). Linear relationship between sulfur and oxygen isotopes from sulfate, like the one observed in our dataset (see Fig. A12) has been evidenced for AOM related processes in estuaries and cold seep environments (Antler et al., 2014). Finding of euhedral Fe-S minerals interpreted as autochthonous organomineralization in sample S17 only (Fig. 5.6 ; see Chapter IV for more details) highlights the enhanced sulfate reduction activity in the interval i6. Additionally, high
151 magnetic susceptibility likely held by greigite (among others, see Ron et al., 2006 ; Fig. A8) and occurrence of framboidal pyrite (Fig. A11), support BSR in the i6 sediment. Oxidation of thermogenic or biologically produced methane by putative ANME communities would allow the release of light carbon in the pore water, leading to an 13C of DIC. Co-occurrence of maxima in S and O isotopes of
13 intense decrease in the ∂ CDIC supports an association of BSR and methanotrophy. 13 sulfate,Minimum and values of minimum observed ∂in the core pore water may even be reflected in C 13 of the aragonite fraction, measured at a the bulkC ∂= -7 ‰ VDPB). This implies precipitation of calciumminimum carbonate in this veryin the same anoxic interval hypolimnion (∂ or in the sediment, as previously suggested in Chapter IV of this manuscript. Calcium carbonate precipitation is indeed favored by AOM through the release of carbonate ions (1), and is regularly observed in methane seeps environments (e. g. Michaelis et al., 2002; Reitner et al., 2005; Pierre and Fouquet, 2007). The microbial communities do not, however, impact the sulfate concentrations. This supports the idea that elemental concentrations in the Dead Sea pore water are largely controlled by the evaporation/precipitation ratio, and the dissolution of evaporitic minerals. Only intense fractionation by microbial activity (such as that induced by sulfate reduction and methanotrophy) may allow singular disturbance of isotopic organic proxies in the Dead Sea pore water.
5.2.6.4 13CDIC 34Ssulfate extrema
AlthoughDiachronous in the same ∂ precipitationand ∂ 34S maximum and
13 C minima are not reached synchronously.dominated period,Albeit poorit is clear resolution, that ∂ this means that AOM∂ occurrence recorded at 64 m cannot solely account for the isotopic signature of sulfate. Potentially, residual sulfate shows similar enrichment in heavy sulfur at this depth, but this interpretation is limited by the sampling resolution. What is clear is that 34 13C is only -9,48 ‰, which is not as depleted as the atoverlying the level values. of maximum Additionally, ∂ S, sample ∂ S18 taken from a similar interval, does not display traces of microbially-influenced Fe-S minerals, indicating that the sediment at this depth did not record exactly similar processes as those of sample S17. AOM may have occurred in this sample, but at smaller scales, as shown by smaller ratio of Ar-OH:Ar. Additionally, the Ar-OH:Ar ratio also suggests a preferential presence of ANME-1 group over ANME-2 and ANME-3 groups for which higher concentrations of Ar-OH were generally
152 documented (Blumenberg et al., 2007). However, this could also be caused by differential degradation of the more labile Ar-OH (Koga et al., 1993; Bouloubassi et al., 2006). Since no DNA could be obtained from this interval (see chapter II, Fig. 2.3), it is anticipated that potential AOM activity in this interval does not occur anymore, and that subsequent degradation may have influenced the Ar-OH:Ar ratio. The whole dataset overall suggest that sulfate reduction either within the AOM framework or as an independent process might have occurred during the whole i6 depositional interval. Its imprint is recorded in sulfur and oxygen isotopes of sulfate. However, AOM by ANME members seems to be the main process triggering the extreme depletion of carbon isotopes as well as the formation of large Fe-S organomineralization, through the enhanced release of HS-. This is seen in sample S17 only.
5.2.6.5 Tentative model for enhanced microbial activity
The reason for such microbial activity is hard to understand, given the poor resolution and lack of information relative to nutrient concentration. During periods of low stand, hypersaline conditions dominate throughout the water column, leaving no suitable environment for the establishment of sustained life. Additionally, the high concentrations of divalent cations in the Dead Sea have often been argued to inhibit microbial development (Oren, 2010a). In turn, periods of lake high-stand do not present such signature either. This is due to the fact that in general, the isotope signature of freshwater inputs masks visualization of such microbial effects. In order to allow for microbes to develop in the Dead Sea realm, and to be active enough to leave their imprint on the geological record we thus need A) a low stand, so that reservoirs are small enough for microbial communities to overprint freshwater signature and B) the establishment a relatively long stratification with significant dilution. In such conditions, sulfate reduction and methanogenesis can establish in anoxic conditions, and influence the geochemistry of the hypolimnion or the pore-water, before they get disturbed by turnover and mixing of the water column. We thus suggest that this specific disturbance at the early Holocene is the result of the recovery from a major lake water decrease before 8.5-8 ka (Migowski et al., 2006), followed by a very humid period, which has allowed the establishment of a diluted epilimnion and development of phytoplanktonic organisms there (as observed by biomarkers). Given the yet unpublished 14C ages (between 8.7 and 7.5 ka), this event can correspond to the humid period evidenced
153 through Soreq cave speleothems during the deposition of sapropel S1 (Orland et al., 2012). Intense decrease of the lake level, together with massive precipitation of gypsum may also have lowered the total divalent concentration of the lake, turning it into a generally more suitable environment for life (Elan Levy, personal communication). In this way, the hypolimnion would host degraders, under the form of fermenters and sulfate reducers. In the shallow sediment, biogenic or thermogenic methane would be oxidized by potential ANME members, leading to the observed trend and lipid profiles. Thermodynamically, AOM is considered to be poorly suited to hypersaline environment such as that of the Dead Sea, as it yields to few energy to allow cells to adjust osmotically (Oren, 2010b). However, AOM has been evidenced in several hypersaline sedimentary environments (Lloyd et al., 2006; Ziegenbalg et al., 2012), indicating that more work has to be pursued in that direction to fully understand this key process.
5.2.7. Conclusion
Microbial activity has been evidenced through combined use of elemental and isotopic profiles of the pore water chemistry, high-resolution lithological characterization and lipid analysis. Anaerobic oxidation of methane is suggested to have disturbed organic proxies at the beginning of the Holocene. The intense water decrease at the end of Pleistocene, and relatively stable meromictic conditions in the lake, during a generally warm period may have allowed the development of a consortium of ANME and SRB. The potential for such microbial disturbance must be taken into account when attempting to reconstruct paleoclimates using O, C and S isotope proxies, as well as magnetic or Fe fractions. Even in extreme environments, the effects of life must not be underestimated, as it is clear that limits for its development, especially in hypersaline environments, are still not established.
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Appendix Material
Fig. A7 Mass spectrum of F2 compound of S17, interpreted as penthamethylicosene with two double bonds (Pmi).
Fig. A8 Partial chromatogram of fraction F3 of samples S17 to S21. Note major peaks of tetrahymanol for S17 and S19. ▼: silylated unbranched alcohol. ● : saturated n- alkanes; Ar: Archaeol. Ext. Ar: extended Archaeol. Poll: artifact from protocole.
Fig. A9 Mass spectra of macrocyclic glycerol di-ether compounds (a-g) and macrocyclic archaeol (h)
Fig. A10 Gregite as small white spots in the upper left corner f the photograph. Sample taken from core 5017-1-a-34a1 (i6 zone, ca. 63 m blf). The large central mineral is titano-magnetite. Note occurrence of euhedrak Fe-S mineralizations. In P6 ziben high magnetic susceptibility is most probably carried by these phases. Photo credits and courtesy of Ute Frank, Norbert Nowaczyk and Ina Neugebauer.
Fig. A11 SEM picture of framboidal pyrite taken from core 5017-1-A-34-1 (i6 zone, ca 63 m blf). Note formation of framboid through aggregation of euhedral iron sulfides, like suggested in Chapter IV of the manuscript. Photo credits and courtesy of Ute Frank, Norbert Nowaczyk and Ina Neugebauer.
Fig. A12 18 34S of pore water sulfate from 58 to 94 m below lake floor (core 5017-1A). Linear relationship of δ O and δ
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162 Chapter VI The subsurface biosphere of the Dead Sea and lacustrine geomicrobiology
“I should live in salt”
Trouble will find me, The National
163 Chapter VI. Conclusion
164 Chapter VI. Conclusion
6.1 Towards the building of a unified model for geomicrobiology in the Dead Sea subsurface
Along this manuscript, we have intended to give a general view of current life in the Dead Sea subsurface, but also of its dynamic and former activity. This was done through the recollection data using varied methods in the core (Fig. 6.1.) and in an active microbial mat of the Dead Sea shore. Several key points have arisen:
1) Current microbial life is known to exist until depths of 90 m below the lake floor of the Dead Sea. DNA was retrieved at even larger depths (200 mblf). This life is dominated by halophilic Archaea of the Halobacteria class. Other groups, known to be well adapted to hypersaline anoxic conditions such as archaeal MSBL1 Candidate Divisions and KB1 Candidate Divisions have been identified based on their 16S rRNA gene fragments. The latter one is among the main clear bacterial contributor identified in the Dead Sea sediment. However, all these groups show relatively low similarities with their database closest matches, indicating that numerous new species and potentially new genera exist in such sediment. The next challenge will then be to properly identify these species, possibly through cultivation of these members. Incorporation of new assembly methods for investigating rare groups from poorly known environments should also complete standard metagenomics techniques.
2) High-throughput sequencing techniques have allowed accessing the metabolic potential of some of the communities inhabiting the Dead Sea sediment. It has been shown that methanogenesis, mainly through the degradation of methylated compounds is performed in an aragonitic level at 2.7 m. Its actors both are members of the archaeal classes Methanobacteria and Methanomicrobia as well as members of the MSBL1 Candidate Division, which may benefit from the degradation of osmotic solutes such as glycine betaine, by members of the KB1 Candidate Division. Additionally, the potential for sulfate reduction and fermentation from numerous substrates is shown. At 90 m deep, members of the Halobacteria class only seem to have adapted to this environment and can live from the fermentation sugar alcohols or amino and organic acids. In both
165 Chapter VI. Conclusion
sediments, the potential for osmotic equilibration by salt-in and salt-out strategies has also been highlighted, widening the classic view of the Dead Sea hypersaline environment. Work towards the recognition of the bacterial potential in those sediments will complete the understanding of microbial interaction in this extreme subsurface environment. Among others, the recycling of the available organic matter, and its potential successive degradational steps should be investigated, to constrain synthrophic and consortium relationship. Work using proteomics or RNA may also bring information on the active pathways at stake in the sediment.
3) Although presence and metabolic potential of microbes in the deep sediment have to a certain extent been defined, little information remains available on the current activity of such microbes in the subsurface. We observed traces of this activity mainly under the form of Fe-S mineralization. Microbial sulfate reduction provides reduced sulfur that reacts with ferrous iron to form Fe-S spherulitic precursors, which can aggregate to form greigite, framboidal clusters as well as euhedral pyrite. Additionally, the finding of remnant of external polymeric substances, arguably produced by microbes in the sediment, proves that microbes do not only exist in the sediment in a dormant stage. Such products serve as carbon sources to sedimentary communities, but can also serve as template for the precipitation of aragonite in the sediment. It suggests that microbes have an influence on the mineralogical record in the Dead Sea subsurface environment. Additional work involving tracing of radioactive carbon incorporation by microbes within mineral lattices in synthetic Dead Sea-like environment should help qualify the extent of such processes, and quantify the effect of microbial life onto the sedimentary record.
4) The combined analysis of pore water isotopes and lipid biomarkers have allowed to establish that the sulfur and carbon cycles have been disturbed, notably during the Holocene, probably by anaerobic oxidation of methane coupled to sulfate reduction. It shows that the Dead Sea biological history is rather varied and dynamic, and that humid periods have allowed the development of planktonic eukaryotes, possibly having an influence on the sedimentary record and its
166 Chapter VI. Conclusion
microbial population, and subsequently disturbing environmental proxy susceptible to be used for paleoclimatic reconstructions.
The integration of these varied dataset into a wider time frame allows the building of a well-supported, yet still hypothetical model for life development in the Dead Sea subsurface (Fig. 6.2). Identified similarities and differences in lithological facies call for the definition of a “microbial facies” during periods of dominant evaporation. It is defined by members of the Halobacteria class, which live in the sediment by fermentation of available substrate, at a supposedly slow growth rate. They are different from identified communities inhabiting aragonite-dominated sediments. In these facies, traces of sulfate reduction, methanogenesis and anaerobic oxidation of methane have been found suggesting a more varied microbial activity. This probably arises from the formation of a diluted epilimnion during humid periods. Nutrient input, together with lower salinity allows the development of halotolerant planktonic communities, which would provide a wide range of substrate to the communities living in the hypersaline brine or brine sediment interface. These communities will then be trapped within the sediment while sedimentation continues. They can keep on subsisting on the available substrate, until it is completely consumed or until exhaustion of nutrients. We hypothesize that although replenishment may occur via fluid circulation or microbial motility, no microbes will further establish in such micro-niches, leaving the aragonitic sediment relatively deprived of extractable DNA, as observed in deeper AAD intervals. Consecutively, current life in the Dead Sea sediment is the result of the establishment of microbial communities at the time of sedimentation. Little to no turnover will occur in such setting, possibly because of poor organic matter quality and extreme salinity implying slow growth rates requiring good adaptation. Additionally, high sedimentation rates would lead to fast burial of these communities. Since the subsurface biosphere derives from that inhabiting the deep brine at time of sedimentation, and that the deep brine communities develop on the substrates made available by production in the upper water column, either in a mixed monolimnion (during arid periods) or in a diluted epilimnion (during humid periods), we suggest that today’s microbial assemblages are influenced by climatic conditions prevailing at time of sedimentation.
167 Chapter VI. Conclusion
Fig. 6.1: Summary of the analysis carried out on the core material within the course of this PhD project. DNA extractions from fluid inclusions were omitted for the sake of clarity. In orange are the samples for which metagenomes were studied.
6.2 Reactions of microbial ecosystems to a changing environment: can the past of lakes be a key to the future?
6.2.1 From surface to bottom: microbial action in a changing environment.
Global net primary production comes for about 50% from phytoplanktonic activity in the oceans (Field et al., 1998). The resulting organic matter is partially degraded in the water column, and exported to the seafloor. Numerous studies have concentrated on the fate of this sedimentary organic matter, as it feeds a rich and diverse microbial community (DeLong, 2004; Schippers et al., 2005; Kallmeyer et al., 2012) and plays a key role in the global carbon cycle. It highlights the importance for the integration of microbial communities study from the water column to the sediment. In times of climate change concern, understanding the ecological variations of these communities is even more important since microbes form the first and last links of the carbon chain.
168 Chapter VI. Conclusion
Fig. 6.2: model for microbial assemblage settlement in the Dead Sea sediment.
However, changes in diversity and activity are hard to account for, as the ocean is a large and diverse system. Additionally, monitoring of these changes and their impact on the whole carbon cycle is complicated by the cost of research operations on the ocean (cruises on a huge surface, and sampling at various depths and sites). Finally, approaching climate changes and its impact on the ocean biota remains difficult due to its complex dynamic and its buffering capacity. In this way, oceans remain relatively stable environments where ecosystem variations can only be observed in long term studies at carefully selected scales.
169 Chapter VI. Conclusion
6.2.2. When subsurface microbiology reflects climatic variations: geomicrobiology of lacustrine environments
Comparatively, and although less important in the global carbon cycle, lakes display advantages that can be determinant for modeling ecosystem reaction to climate changes. They are more susceptible to be affected by climate variations due to their smaller volume. Their changes are also faster and sometimes observable at the scale of research project durations. Finally, they are generally more accessible and their diversity in size, physico-chemical conditions and geographical setting can be exploited to address questions in environments of pinpointed interest. Recently, the study of subsurface microbial communities in lakes has revealed the prevalent influence of climatic conditions on their metabolic distribution. Indeed, works in Laguna Potrok Aike, Argentina (Vuillemin et al., 2013) , and in Lake Van, Turkey (Glombitza et al., 2013), have emphasized the importance of substrate type, generally determined by climate variations, in the establishment of peculiar microbial assemblages in the sediment. The geomicrobiology study of the Dead Sea sediment tends to show similar trends, with specificities owing to its extreme salinity. The Dead Sea is indeed subject to intense changes induced both by climate and human activity. Now, its level is dramatically decreasing primarily because of 1) the use of its tributary water for agricultural purpose, and 2) fastened evaporation for potash and magnesium chloride extraction by Dead Sea Works and Arab Potash companies (Tahal Group and GSI, 2011). In this context, research in aquatic microbiology has allowed the characterization of extreme halophilic life and established a good view of its extent in the Dead Sea water column (e.g. Oren and Ventosa, 1999; Oren, 2010; Oren and Gunde-Cimerman, 2012). Dynamics of its extreme populations under these changing conditions are also better constrained (Oren, 1993; Oren et al., 1995; Bodaker et al., 2010; Rhodes et al., 2012). The results obtained in the course of this project suggest a peculiar dynamic in the sediment. Although selected by the extreme salinity, microbes identified in the sediment seem to be primarily influenced by the lake conditions they have been buried under. In aragonitic sediments, typical of periods of lake stratification and upper layer dilution, communities form a rather complete trophic chain, involving fermenting, sulfate- reducing and methanogenic halophiles. These communities, probably developing in the
170 Chapter VI. Conclusion anoxic hypersaline hypolimnion, are hypothesized to remain active upon burial in the sediment, until they run out of substrate. Alternatively, in halite-gypsum sediments characteristic of the monomictic stage, Archaea of the Halobacteria class seem to be the only organisms able to cope with the harshness of an undiluted water column (Bodaker et al., 2010). They are found in sediments down to 200 m depth, suggesting little turnover and thus very slow activity in these facies. Thus, in the Dead Sea, microbial assemblages are related to physico-chemical parameters of the lake. The latter being directly induced by climatic variations, sedimentary communities are selected by the climatic conditions undergoing in the Levantine area at the time of sedimentation. The suggestive results obtained from the Dead Sea deep subsurface case study demonstrate the potential of multidisciplinary approaches for unraveling ecosystem changes in lacustrine environment, through the use of deep scientific drilling.
6.2.3. Collaborative approach for understanding future changes in the aquatic biosphere of lakes and oceans.
The extent to which microbial ecosystems can react and be affected by climatic changes can be approached by integrated multidisciplinary studies. One of the final goals is to construct models addressing changes in microbial ecology and its putative implications for marine biology and the whole carbon cycle in general. By focusing on contrasted environments such as lakes, we chose highly dynamic subjects, with in most cases easy access to sampling spots. A view of past changes is provided by the addition of subsurface studies. The recent findings from the Dead Sea suggest that microbial communities in the lake water column are relatively similar to those existing during the last interglacial period that is 70 to 80 ka ago. These conclusions directly depend on the monitoring of present communities. Finally, biogeochemical studies integrated to the whole water column and sediment domain allows completing the path of carbon, until its recycling as CO2 or methane, or its accumulation through sedimentary organic matter. The idea behind this final section is best conveyed by figure 6.3, encompassing common research themes of environmental microbiology and oceanography labs, and adding subsurface microbiology and biogeochemistry in lacustrine settings, in order to better predict the “ecosystems evolution and reaction” to a changing environment.
171 Chapter VI. Conclusion
Being either in water-stressed regions (e.g. Dead Sea, Levant), densely populated places (e.g. Great Lakes in North America or in Africa), or pristine areas (e.g. Potrok Aike in Patagonia or El’gygytgyn in Siberia), lakes allow the study of relevant environments for climate change studies. They also are relatively small ecosystems, with reasonably reachable ins and outs. A final objective is thus to clarify the importance of such multidisciplinary approach in the study of climate impact on microbial ecosystems. This could be done by building a checklist of collaborative domains and methods susceptible to help characterizing the dynamics of microbial communities in water. The lake toolbox can be further proposed (and ideally adapted) for application to microbial oceanography in order to better constrain the “ecosystems evolution and reaction” target most environmental studies are aiming for. Integration of various scientific backgrounds, related to aquatic sciences will help reaching this goal.
Fig. 6.3: Collaborative pathway for the study of ecosystems evolution and reaction in a changing environment.
172 Chapter VI. Conclusion
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174 Appendix
175
176 Part II. Table A1. OTU de inition and presence/absence in each sample.
OTU Phylum Class Order Family Genus Identity (%) CT1 CT2 CT3 CT4 M2 M1
� ARCHAEA 30 25 31 23 15 20
KM587793 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halobacterium 95.19 1 0 0 0 0 0
KM587812 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halomicrobium 95.2 1 0 0 0 0 0
KM587805 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halomicrobium 88.82 1 0 0 0 0 0
KM587811 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halomicrobium 70.41 1 0 0 0 0 0
KM587753 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halonotius 96.2 0 0 1 0 0 0
KM587781 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halonotius 91.45 0 0 0 0 0 1
KM587831 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halonotius 91.4 0 0 0 1 0 0
KM587774 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halonotius 91.25 0 0 1 0 0 0
KM587773 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halonotius 87 0 0 1 0 0 0
KM587799 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Haloplanus 91 1 0 0 0 0 0
KM587748 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 97.8 1 0 1 1 1 1
KM587824 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 97 0 0 0 1 0 0
KM587827 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 96.4 0 0 1 1 0 0
KM587784 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 95.6 0 0 0 0 0 1
KM587776 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 95.4 0 0 1 0 0 0
KM587818 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 95.4 1 0 0 0 0 0
KM587832 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 95.4 0 0 0 1 0 0
KM587821 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 95.2 0 0 0 1 0 0
KM587779 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 95 0 0 0 0 0 1
KM587822 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 94.8 0 0 1 1 1 0
KM587810 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 94.8 1 0 0 0 0 0
KM587833 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 94.8 0 0 0 1 0 0
KM587755 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 94.6 0 0 1 1 1 0
KM587820 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 94.6 0 0 0 1 0 0
KM587803 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 94.4 1 0 0 0 0 0
A 11_11 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 94.2 0 0 0 1 0 0
KM587827 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 93.81 0 0 0 1 0 0
KM587763 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 93.8 0 0 1 0 0 0
KM587770 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 93.8 0 0 1 0 0 0
KM587792 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 93.8 1 0 0 0 0 0
KM587814 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 93.21 1 0 0 0 0 0
KM587830 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 93.2 0 0 0 1 0 0
KM587780 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 93.01 0 0 0 0 0 1
KM587756 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 93 0 0 1 0 0 0
KM587767 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.81 0 0 1 0 0 0
KM587771 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.81 0 0 1 0 0 0
KM587794 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.8 1 0 0 0 0 0
KM587819 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.61 0 0 0 1 0 0
KM587825 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.61 0 0 0 1 0 0
KM587775 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.6 0 0 1 0 0 0
KM587804 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.56 1 0 0 0 0 0
KM587754 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.4 0 0 1 0 0 0
KM587782 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.4 0 0 0 0 0 1
KM587817 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92.22 1 0 0 0 0 0
KM587758 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 92 0 0 1 0 0 1
KM587813 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 91.85 1 0 0 0 0 0
KM587762 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 91.62 0 0 1 0 0 0
KM587764 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 91.6 0 0 1 0 0 0
KM587801 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 91.4 1 0 0 0 0 0
KM587765 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 91.2 0 0 1 1 0 1
KM587786 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90.98 0 0 0 0 1 0
KM587769 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90.6 0 0 1 0 0 0
KM587761 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90.42 0 0 1 0 0 1
KM587766 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90.42 0 0 1 0 0 0
KM587777 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90.42 0 0 1 0 0 0
KM587783 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90.4 0 0 0 0 0 1
KM587800 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90.4 1 0 0 0 0 0
KM587760 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90.2 0 0 1 0 0 0
KM587826 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 90 0 0 0 1 0 0
KM587789 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 89.26 0 0 0 0 1 0
KM587791 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 89.24 1 0 0 0 0 0
KM587806 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 89.24 1 0 0 0 0 0
177 KM587772 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 89.22 0 0 1 0 0 0
KM587829 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 89.2 0 0 0 1 0 0
KM587750 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 89.07 0 0 1 0 0 0
KM587752 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 88.47 0 0 1 0 0 0
KM587793 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 88.25 1 0 0 0 0 0
KM587798 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 87.94 1 0 0 0 0 0
KM587828 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 87.8 0 0 0 1 0 0
KM587816 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 87.74 1 0 0 0 0 0
KM587768 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 87.65 0 0 1 0 0 0
KM587757 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 87.6 0 0 1 0 0 0
KM587822 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 86.65 0 0 0 1 0 0
KM587823 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 86 0 0 0 1 0 0
KM587809 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 83 1 0 0 0 0 0
KM587797 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 82.42 1 0 0 0 0 0
KM587785 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 80 0 0 0 0 0 1
KM587835 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 78.33 0 0 0 0 0 1
KM587808 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 71.37 1 0 0 0 0 0
KM587807 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Haloterrigena 82.64 1 0 0 0 0 0
KM587751 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae uncultured 94.13 1 0 1 1 1 1
KM587749 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae uncultured 94 0 0 1 0 0 0
KM587834 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Uncultured 92.22 0 0 0 1 0 0
KM587796 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Uncultured 91.62 1 0 0 0 0 0
KM587726 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae uncultured 88.78 0 1 0 0 0 0
KM587802 Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Uncultured 82.91 1 0 0 0 0 0
KM587815 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 90.2 1 0 0 0 0 0
KM587884 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 89.8 0 0 0 0 1 0
KM587843 Euryarchaeota Halobacteria Halobacteriales MSP41-clade uncultured 89.4 0 0 0 0 0 1
KM587841 Euryarchaeota Halobacteria Halobacteriales MSP41-clade uncultured 88.8 0 0 0 0 0 1
KM587887 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 88.4 0 0 0 0 1 0
KM587845 Euryarchaeota Halobacteria Halobacteriales MSP41-clade uncultured 87.03 0 0 0 0 0 1
KM587842 Euryarchaeota Halobacteria Halobacteriales MSP41-clade uncultured 85.6 0 0 0 0 0 1
KM587778 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 83.76 0 0 0 0 0 1
KM587790 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 81.4 0 0 0 0 1 0
KM587882 Euryarchaeota Halobacteria Halobacteriales MSP41-clade uncultured 81.11 0 0 0 0 1 0
KM587886 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 81.04 0 0 0 0 1 0
KM587846 Euryarchaeota Halobacteria Halobacteriales MSP41-clade uncultured 80.36 0 0 0 0 0 1
KM587883 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 80.28 0 0 0 0 1 0
KM587787 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 79.8 0 0 0 0 1 0
KM587885 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 76.23 0 0 0 0 1 0
KM587788 Euryarchaeota Halobacteria Halobacteriales MSP41-clade Uncultured 76.1 0 0 0 0 1 0
KM587844 Euryarchaeota Halobacteria Halobacteriales MSP41-clade uncultured 72.15 0 0 0 0 0 1
KM587739 Euryarchaeota Methanomicrobia Unknown ST-12K10A Uncultured 90.32 0 1 0 0 0 0 Candidate Division KM587723 Euryarchaeota Thermoplasmata Uncultured 85.2 0 1 0 0 0 0 MSBL1 Candidate Division KM587744 Euryarchaeota Thermoplasmata Uncultured 85 0 1 0 0 0 0 MSBL1 Candidate Division KM587724 Euryarchaeota Thermoplasmata Uncultured 84.43 0 1 0 0 0 0 MSBL1 Candidate Division KM587734 Euryarchaeota Thermoplasmata Uncultured 83.95 0 1 0 0 0 0 MSBL1 Candidate Division KM587732 Euryarchaeota Thermoplasmata Uncultured 83.83 0 1 0 0 0 0 MSBL1 Candidate Division KM587741 Euryarchaeota Thermoplasmata Uncultured 83.4 0 1 0 0 0 0 MSBL1 Candidate Division KM587729 Euryarchaeota Thermoplasmata Uncultured 83.23 0 1 0 0 0 0 MSBL1 Candidate Division KM587738 Euryarchaeota Thermoplasmata Uncultured 83.23 0 1 0 0 0 0 MSBL1 Candidate Division KM587736 Euryarchaeota Thermoplasmata Uncultured 82.83 0 1 0 0 0 0 MSBL1 Candidate Division KM587728 Euryarchaeota Thermoplasmata Uncultured 82.57 0 1 0 0 0 0 MSBL1 Candidate Division KM587731 Euryarchaeota Thermoplasmata Uncultured 82.55 0 1 0 0 0 0 MSBL1 Candidate Division KM587737 Euryarchaeota Thermoplasmata Uncultured 82.55 0 1 0 0 0 0 MSBL1 Candidate Division KM587730 Euryarchaeota Thermoplasmata Uncultured 82.51 0 1 0 0 0 0 MSBL1 Candidate Division KM587742 Euryarchaeota Thermoplasmata Uncultured 81.88 0 1 0 0 0 0 MSBL1 Candidate Division KM587747 Euryarchaeota Thermoplasmata Uncultured 81.1 0 1 0 0 0 0 MSBL1 Candidate Division KM587746 Euryarchaeota Thermoplasmata Uncultured 80.71 0 1 0 0 0 0 MSBL1 Candidate Division KM587725 Euryarchaeota Thermoplasmata Uncultured 79.8 0 1 0 0 0 0 MSBL1 Candidate Division KM587727 Euryarchaeota Thermoplasmata Uncultured 79.8 0 1 0 0 0 0 MSBL1 Candidate Division KM587740 Euryarchaeota Thermoplasmata Uncultured 76.62 0 1 0 0 0 0 MSBL1 Candidate Division KM587735 Euryarchaeota Thermoplasmata Uncultured 76.23 0 1 0 0 0 0 MSBL1 Candidate Division KM587743 Euryarchaeota Thermoplasmata Uncultured 76.08 0 1 0 0 0 0 MSBL1 Candidate Division KM587733 Euryarchaeota Thermoplasmata Uncultured 75.98 0 1 0 0 0 0 MSBL1 Candidate Division KM587745 Euryarchaeota Thermoplasmata Uncultured 73.08 0 1 0 0 0 0 MSBL1
178 BACTERIA 3 15 0 0 23 Candidate Division KM587868 Uncultured 99 0 1 0 0 0 KB1 Candidate Division KM587840 Uncultured 98.4 0 1 0 0 1 KB1 Candidate Division KM587857 Uncultured 97.4 0 0 0 0 1 KB1 Candidate Division KM587849 Uncultured 97 0 0 0 0 1 KB1 Candidate Division KM587852 Uncultured 96.61 0 0 0 0 1 KB1 Candidate Division KM587866 Uncultured 96.44 0 1 0 0 0 KB1 Candidate Division KM587854 Uncultured 96.41 0 0 0 0 1 KB1 Candidate Division KM587871 Uncultured 94.2 0 1 0 0 0 KB1 Candidate Division KM587879 Uncultured 94.02 0 0 0 0 1 KB1 Candidate Division KM587836 Uncultured 93.61 0 0 0 0 1 KB1 Candidate Division KM587858 Uncultured 93.61 0 0 0 0 1 KB1 Candidate Division KM587853 Uncultured 93.6 0 0 0 0 1 KB1 Candidate Division KM587859 Uncultured 93.6 0 0 0 0 1 KB1 Candidate Division KM587850 Uncultured 92.83 0 0 0 0 1 KB1 Candidate Division KM587872 Uncultured 92.81 0 1 0 0 0 KB1 Candidate Division KM587837 Uncultured 92.61 0 0 0 0 1 KB1 Candidate Division KM587864 Uncultured 92.42 0 1 0 0 0 KB1 Candidate Division KM587874 Uncultured 90.6 0 1 0 0 0 KB1 Candidate Division KM587865 Uncultured 89.64 0 1 0 0 0 KB1 Candidate Division KM587873 Uncultured 87.08 0 1 0 0 0 KB1 Candidate Division KM587875 Uncultured 84.71 0 1 0 0 0 KB1 Candidate Division KM587870 Uncultured 82.63 0 1 0 0 0 KB1 Candidate Division KM587847 Uncultured 80.12 0 0 0 0 1 KB1 Candidate Division KM587869 Uncultured 72 0 1 0 0 0 KB1 Candidate Division KM587862 Uncultured 68 0 0 0 0 1 KB1 KM587867 Deferribacteres Deferribacteres Deferribacterales SAR406 Clade (Marine Group A) Uncultured 75.84 0 1 0 0 0
KM587876 Deferribacteres Deferribacteres Deferribacterales SAR406 Clade (Marine Group A) Uncultured 75.05 0 1 0 0 0
KM587863 Deferribacteres Deferribacteres Deferribacterales SAR406 Clade (Marine Group A) Uncultured 73.86 0 1 0 0 0 Deinococcus- KM587877 Deinococci Deinococcales Trueperaceae Truepera 91.02 1 0 0 0 0 Thermus KM587855 Firmicutes Clostridia Clostridiales Halanaerobiaceae Halanaerobium 89.6 0 0 0 0 1
KM587856 Firmicutes Clostridia Clostridiales Halanaerobiaceae Halanaerobium 89.4 0 0 0 0 1
KM587838 Firmicutes Clostridia Clostridiales Halanaerobiaceae Halanaerobium 89 0 0 0 0 1
KM587860 Firmicutes Clostridia Clostridiales Halanaerobiaceae Halanaerobium 88.4 0 0 0 0 1
KM587839 Firmicutes Clostridia Clostridiales Halanaerobiaceae Halanaerobium 86.83 0 0 0 0 1
KM587880 Firmicutes Clostridia Clostridiales Halanaerobiaceae Halanaerobium 86.6 0 0 0 0 1
KM587881 Nitrospirae Nitrospira Nitrospirales Nitrospiraceae Nitrospira 86.86 1 0 0 0 0
KM587861 Nitrospirae Nitrospira Nitrospirales Nitrospiraceae Uncultured 80.04 0 0 0 0 1
KM587878 Proteobacteria Deltaproteobacteria Desulfobacterales Nitrospinaceae Uncultured 81.78 1 0 0 0 0
KM587861 Proteobacteria Deltaproteobacteria Desulfovibrionales Desulfohalobiaceae Desulfovermiculus 86.51 0 0 0 0 1
KM587848 Proteobacteria Gammaproteobacteria Order Incertae Sedis Family Incertae Sedis Thiohalorhabdus 85.93 0 0 0 0 1
179 Table A2. OTU distribution and phylotypes de�inition DNA1 COA2 DNA32 DS57 CT9 CT8 Domain Phylum Class Order Family Genus surface halite aad 2.74 m gypsum 90.64 m halite 206.53 m aragonite mat halite-gypsum mat Archaea Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halobacterium 1 0 0 0 0 0 Archaea Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halomicrobium 2 0 0 0 0 0 Archaea Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halonotius 0 0 3 1 0 1 Archaea Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Haloplanus 1 0 0 0 0 0 Archaea Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Halorhabdus 19 0 26 20 5 11 Archaea Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae Haloterrigena 1 0 0 0 0 0 Archaea Euryarchaeota Halobacteria Halobacteriales Halobacteriaceae uncultured 3 1 2 2 1 1 Archaea Euryarchaeota Halobacteria Halobacteriales MSP41-clade uncultured 1 0 0 0 9 7 Archaea Euryarchaeota Methanomicrobia uncultured 0 1 0 0 0 0 Archaea Euryarchaeota Thermoplasmata MSBL1 Candidate Division uncultured 0 23 0 0 0 0 Archaea unknown 1 1 0 0 0 0 0 Archaea unknown 2 1 0 0 0 0 0 TOTAL 30 25 31 23 15 20
Bacteria Deferribacteres Deferribacteres Defferibacterales SAR406 Clade (Marine Group A) uncultured 0 3 0 0 0 Bacteria Deinococcus-Thermus Deinococci Deinococcales Trueperaceae Truepera 1 0 0 0 0 Bacteria Firmicutes Clostridia Halanaerobiales Halanaerobiaceae Halanaerobium 0 0 0 0 6 Bacteria KB1 Candidate Division uncultured 0 11 0 0 13 Bacteria Nitrospirae Nitrospira Nitrospirales Nitrospiraceae Nitrospira 1 0 0 0 0 Bacteria Nitrospirae Nitrospira Nitrospirales Nitrospiraceae uncultured 0 0 0 0 1 Bacteria Proteobacteria Deltaproteobacteria Desulfobacterales Nitrospinaceae uncultured 1 0 0 0 0 Bacteria Proteobacteria Deltaproteobacteria Desulfovibrionales Desulfohalobiaceae Desulfovermiculus 0 0 0 0 1 Bacteria Proteobacteria Gammaproteobacteria Order Incertae Sedis Family Incertae Sedis Thiohalorhabdus 0 0 0 0 1 Bacteria unknown 1 0 1 0 0 Bacteria unknown 2 0 0 0 0 1 TOTAL 3 15 0 0 23
Table A3. Calibration values for microthermometry analysis
sample t° min t°f Th t° theoretical t° measured - 0 320 0 0.0 - 0 310.5 374 381.0 std H2O (D509-061) - 0 310.2 10 10.1 - 0 310.3 -56.6 -56.5 - - 382.5 std H2O (93-0) - - 380.1 Calibration line equation - - 380.5 x= (y-0,3673)/1,0175 -56.5 10 - std H2O-CO2 -56.4 10.1 - -56.5 10.1 -
180 Table A4. Temperatures of homogenization for 4 samples of the Holocene part of the core
sample Th Th corrected 27.1 26.3 25.8 25.0 22.1 21.4 20.2 19.5 21.1 20.4 21.4 20.7 25.4 24.6 19.5 18.8 A-31-A1 - 59.61 m blf 19.5 18.8 18.7 18.0 23.5 22.7 22 21.3 21.5 20.8 26.8 26.0 23.4 22.6 24 23.2 20.4 19.7 25.4 24.6 25.4 24.6 25.1 24.3 27.8 27.0 27 26.2 26.5 25.7 26.4 25.6 25.5 24.7 23.4 22.6 24.6 23.8 23.5 22.7 25.5 24.7 25.4 24.6 1-H-1-CC - 0.24 m blf 25.2 24.4 35 34.0 31.8 30.9 26.1 25.3 24.5 23.7 29.8 28.9 29.6 28.7 29.9 29.0 24.8 24.0 25.5 24.7 25.7 24.9 25.8 25.0 24.9 24.1 32.5 31.6 32 31.1 28.9 28.0 DNA17 28.5 27.6 28 27.2 30 29.1 29.4 28.5 50 48.8 DNA19 48.7 47.5 51.8 50.5
181 Part II Fig. A1 Photograph of sediment sample incubated with tyramide (CARD-FISH dye) and showing non nt speci�ic binding of the dye (green) onto the sedime
Fig. A2 16S rRNA gene based phylogenetic tree of Bacteria representative sequences for phylotypes estimated difference in nucleotide sequences. bars on the right panel identify main bacterial classes de�ined in this study. tree was built using the Waighted Neighbor Joining method.5 µm Scale 10% of and major clustersDEAD SEA CT2 identi�iableKM587867 in the Dead Sea sediment 100 DEAD SEA CT2 KM587876 uncultured Deferribacterales 70 DEAD SEA CT2 KM587863 GQ860143 Nitrospira Ohio river sediment 100 DEAD SEA CT1 KM587881 Nitrospira DEAD SEA CT1 KM587878 100 EU134581 unc Nitrospinaceae Oklahoma prairie soil
76 FN393488 Desulfovermiculus Tunisian solar saltern δ-proteobacteria 61 DEAD SEA M2 KM587851 100 FJ536441 Desulfovermiculus Mediterranean solar saltern sediment AB637080 Truepera Japanese soil 94 DEAD SEA CT1 KM587877 Deinococci
66 EU687421 Thiohalorhabdus Mexican sabkha microbial mat 100 DEAD SEA M2 KM587848 γ-proteobacteria
DEAD SEA M2 KM587856 DEAD SEA M2 KM587860 65 EU245295 Halanaerobium Puerto Rican hypersaline lagoon mat 54 DEAD SEA M2 KM587838 94 Clostridia 99 DEAD SEA M2 KM587855 98 DQ103656 Halanaerobium Israeli endoevaporitic microbial mat DEAD SEA M2 KM587839 84 DEAD SEA M2 KM587880 100 100 EU245123 Halanaerobium Puerto Rican hypersaline lagoon mat
56
60 HQ425220 Candidate Division KB1 unc Iranian salt lake AJ133617 Candidate Division KB1 unc Kebrit Deep brine water DEAD SEA CT2 KM587871 DEAD SEA M2 KM587852 DEAD SEA CT2 KM587868 DEAD SEA CT2 KM587840
50 DEAD SEA M2 KM587854 DEAD SEA M2 KM587879 DEAD SEA CT2 KM587873 100 50 DEAD SEA CT2 KM587865 DEAD SEA M2 KM587862 DEAD SEA M2 KM587849 DEAD SEA M2 KM587858 DEAD SEA M2 KM587847 KB1 Candidate Division DEAD SEA M2 KM587836 DEAD SEA M2 KM587850 DEAD SEA CT2 KM587875 DEAD SEA M2 KM587857 DEAD SEA M2 KM587859 DEAD SEA M2 KM587853 DEAD SEA M2 KM587837 HQ425219 Candidate Division KB1 unc Iranian salt lake EF106489 Candidate Division KB1 unc Mexican salt crust DEAD SEA CT2 KM587874 DEAD SEA CT2 KM587870 DEAD SEA CT2 KM587872
53 DEAD SEA CT2 KM587864 AJ133615 Candidate Division KB1 unc Kebrit Deep brine water
82 DEAD SEA CT KM587866 73 DEAD SEA CT2 KM587869
182 Scale: 0.1 Fig. A3 Comparison of microbial diversity in sample CT2 and CT3 obtained through 16S rRNA cloning (this study) and metagenomic data (A) Microbial diversity in the aragonitic sample. Metagenomes data from assem- bled 16S rRNA genes obtained from AD metagenome presented in Thomas et al. (2014). (B) Comparison of archaeal diversity. Since metagenomic data did not provide convincing 16S rRNA gene fragments for sample GY A(Thomas et al., 2014), comparison is made through total gene similarity Bwith the M5NR database. 100% 100% 90% 90% Other Bacteria 80% Euryarchaeota 80% Proteobacteria 70% Archaea 70% Gammaproteobacteria Deltaproteobacteria 60% Halobacteria 60% Methanothermobacter Deferribacterales 50% 50% Methanomicrobia Desulforvermiculus 40% 40% Methanomicrobia ST12K10A KB1 Candidate Division 30% Thermoplasmata 30% 20% MSBL1 candidate Division 20%
10% 10%
0% 0% Cloned OTUs metagenome Cloned OTUs metagenome Cloned OTUs metagenome (this study) (this study) (this study)
aragonite sample (2.74 m) gypsum sample (90.64 m) aragonite sample (2.74 m)
Fig. A4 Distribution oftemperatures of homogenization of halite �luid inclusions at three different depths of the core. Y-axis is number n of inclusions measured. X-axis is temperature of homogenization Th in °C. At the early Holocene, relatively mild temperature (maximum Th of 26.3 °C and maximum n for 20.5-21.4°C. It seems that surface temperatures of the Dead Sea attained temperatures between 27 and 32 °C in the middle of the Holo- cene, while temperatures recorded in the most recent halite are centered around 24-25 °C. In October 2014, surface water temperatures are �luctuating between 29 and 31°C but are varying widely along the year (http://isramar.ocean.org.il/isramar2009/DeadSea/Default.aspx) 0.24 m 32.9 m 59.61 m 12 3 5
10 4
8 2 3
6
2 4 1
1 2
0 0 0
F
ig. A5 Halite block precipitated during January-February 2010, at around 2 m below the lake surface. This sample is currently being used for calibration of Th and better understanding daily to seasonal temperature variations recorded in the �luid inclusions.
183 Part IV.
Fig. A6 Carbon isotopes of aragonite, detritus and aragonite mixed with detritus laminae for the microbial mat samples at En Qedem. Corresponding TOC is also presented in the table with obtained ∂13CCaCO3
Part V.
Fig. A7 Mass spectrum of F2 compound of S17, interpreted as pentamethylicosene with two double bonds, and of hypothetical non isoprenoidal macrocyclic glycerol diethers (mGD) obtained from sample S17 F4 at t= 58 (mGD 1) and T=61,5 (mGD 2).
184
Fig. A8 Partial chromatogram of fraction 3 of samples S17 to S21. Note major peaks of tetrahymanol for S17 and S19. ▼: silylated unbranched alcohols. ●: saturated n-alkane. Ar: Archaeol. Ext Ar : extended Archaeol. Poll: pollution.
185 Fig. A9 Mass spectra of compounds of mgd compounds (a-g) and macroarchaeol (h).
186 Fig. A10 Greigite as small white spots in the upper left corner of the photograph. Sample taken from core 5017-1-A-34-1 (i6 zone, ca. 63 m blf). The large central mineral is Ti-magnetite. Note occurrence of euhedral Fe-S mineralizations. In P6 zone, high magnetic susceptibility is most probably carried by these phases. Photo credits and courtesy of Ute Frank, Robert Nowaczyk and Ina Neugebauer
Fig. A11 SEM picture of framboidal pyrite taken from core 5017-1-A-34-1 (i6 zone, ca 63 m blf). Note formation of framboid through aggregation of euhedral iron sulfides, like suggested in Part IV. of the manuscript. Photo credits and courtesy of Ute Frank, Robert Nowaczyk and Ina Neugebauer
187 Fig. A12 Linear relationship of δ18O vs δ34S of pore water sulfate from 58 to 94 m below lake floor (core 5017-1A).
25,00 R² = 0,93423 20,00
(‰) 15,00
sulfate 10,00 O 18
δ 5,00
0,00 0,00 5,00 10,00 15,00 20,00 25,00 30,00 35,00 40,00 45,00
34 δ S sulfate (‰)
188